Physics Wiki - User contributions [en]

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2013-04-01T11:13:48Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), MOPS buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
[[File:Nitella_ligature.png|200px|right]]
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming (ligature experiments by Varley in 1844 are shown in the watercolor [right]). What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://eol.org/pages/11583/overview)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie (starring Chara in the culture tank in your BPHS4090 teaching lab). The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

<h1> Appendix Three: APW (artificial pond water) Make-Up </h1>

<p>Make-Up of APW (artificial pond water)</p>

<p>Artificial pond water (APW)
is used to mimic the concentrations of major ions normally found in
freshwater environments, where algae typically grow. The APW stocks are stored
as X100 concentrated solutions in a refrigerator. When diluted to final
concentration, the pH is adjusted to the appropriate level with NaOH. APW 5 is
adjusted to pH 5.0, APW 6 is adjusted to pH 6.0, etc.</p>

<p>X100 stocks are made-up as 100 mM final:</p>

<table border=1 width=600>
<tr><td><b>Salts</b></td><td>Molecular Weight</td><td>grams/100 ml</td>
</tr>
<tr><td>Potassium chloride (KCl)</td><td>74.56</td><td>0.746</td>
</tr>
<tr><td>Calcium chloride, dihydrate (CaCl<sub>2</sub> 2H<sub>2</sub>O)</td><td>147.02</td><td>1.470</td>
</tr>
<tr><td>Magnesium chloride, hexahydrate (MgCl<sub>2</sub> 6H<sub>2</sub>O)</td><td>203.31</td><td>2.033</td>
</tr>
<tr><td>Sodium chloride (NaCl)</td><td>58.44</td><td>0.584</td>
</tr>
<tr><td>Sodium sulfate (Na<sub>2</sub>SO<sub>4</sub>) (for APW10)</td><td>142.04</td><td>1.420</td>
</tr>
<tr><td><b>Buffers</b></td><td></td><td></td>
</tr>
<tr><td>MES (pK<sub>a</sub> 6.15)</td><td>195.23</td><td>1.952</td>
</tr>
<tr><td>MOPS (pK<sub>a</sub> 7.20) </td><td>209.26</td><td>2.092</td>
</tr>
<tr><td>TES (pK<sub>a</sub> 7.50)</td><td>229.25</td><td>2.293</td>
</tr>
<tr><td>TAPS (pK<sub>a</sub> 8.40)</td><td>243.20</td><td>2.432</td>
</tr>
<tr><td>CAPS (pK<sub>a</sub> 10.40)</td><td>221.32</td><td>2.213</td>
</tr>
</table>

<p>For final solutions (final volume 1 liter)</p>

<table border=1 width=600>
<tr><td>100 mM Stock</td><td>KCl</td><td>CaCl<sub>2</sub></td><td>MgCl<sub>2</sub></td><td>NaCl</td><td>Buffer</td>
</tr>
<tr><td><p></p></td><td></td><td></td><td></td><td></td><td align=right>(in ml/1000 ml)</td>
</tr>
<tr><td>APW 5</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 6</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 7</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MOPS</td>
</tr>
<tr><td>APW 8</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TES</td>
</tr>
<tr><td>APW 9</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TAPS</td>
</tr>
<tr><td>APW 10</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 CAPS plus 0.05 Na<sub>2</sub>SO<sub>4</sub></td>
</tr>
</table>

<p>pH is adjusted to final value with NaOH (1 N)</p>

<hr align=left size=1 width="33%">

<p><b>Source:</b> Spanswick RM (1972) Evidence for an electrogenic ion pump in <i>Nitella translucens</i>. I. The effects of pH, K<sup>+</sup>, Na<sup>+</sup>,
light and temperature on the membrane potential and resistence. Biochimica
Biophysica Acta 288:74-89</p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2013-04-01T11:12:42Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), MOPS buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
[[File:Nitella_ligature.png|200px|right]]
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming (ligature experiments by Varley in 1844 are shown in the watercolor [right]). What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://eol.org/pages/11583/overview)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie (starring Chara in the culture tank in your BPHS4090 teaching lab). The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

<h1> Appendix Three: APW (artificial pond water) Make-Up </h1>

<p>Make-Up of APW (artificial pond water)</p>

<p>Artificial pond water (APW)
is used to mimic the concentrations of major ions normally found in
freshwater environments, where algae typically grow. The APW stocks are stored
as X100 concentrated solutions in a refrigerator. When diluted to final
concentration, the pH is adjusted to the appropriate level with NaOH. APW 5 is
adjusted to pH 5.0, APW 6 is adjusted to pH 6.0, etc.</p>

<p>X100 stocks are made-up as 100 mM final:</p>

<table border=1 width=600>
<tr><td><b>Salts</b></td><td>Molecular Weight</td><td>grams/100 ml</td>
</tr>
<tr><td>Potassium chloride (KCl)</td><td>74.56</td><td>0.746</td>
</tr>
<tr><td>Calcium chloride, dihydrate (CaCl<sub>2</sub> 2H<sub>2</sub>O)</td><td>147.02</td><td>1.470</td>
</tr>
<tr><td>Magnesium chloride, hexahydrate (MgCl<sub>2</sub> 6H<sub>2</sub>O)</td><td>203.31</td><td>2.033</td>
</tr>
<tr><td>Sodium chloride (NaCl)</td><td>58.44</td><td>0.584</td>
</tr>
<tr><td>Sodium sulfate (Na<sub>2</sub>SO<sub>4</sub>) (for APW10)</td><td>142.04</td><td>1.420</td>
</tr>
<tr><td><b>Buffers</b></td><td></td><td></td>
</tr>
<tr><td>MES (pK<sub>a</sub> 6.15)</td><td>195.23</td><td>1.952</td>
</tr>
<tr><td>MOPS (pK<sub>a</sub> 7.20) </td><td>209.26</td><td>2.092</td>
</tr>
<tr><td>TES (pK<sub>a</sub> 7.50)</td><td>229.25</td><td>2.293</td>
</tr>
<tr><td>TAPS (pK<sub>a</sub> 8.40)</td><td>243.20</td><td>2.432</td>
</tr>
<tr><td>CAPS (pK<sub>a</sub> 10.40)</td><td>221.32</td><td>2.213</td>
</tr>
</table>

<p>For final solutions (final volume 1 liter)</p>

<table border=1 width=600>
<tr><td>100 mM Stock</td><td>KCl</td><td>CaCl<sub>2</sub></td><td>MgCl<sub>2</sub></td><td>NaCl</td><td>Buffer</td>
</tr>
<tr><td><p></p></td><td></td><td></td><td></td><td></td><td align=right>(in ml/1000 ml)</td>
</tr>
<tr><td>APW 5</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 6</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 7</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MOPS</td>
</tr>
<tr><td>APW 8</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TES</td>
</tr>
<tr><td>APW 9</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TAPS</td>
</tr>
<tr><td>APW 10</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 CAPS plus 0.05 Na<sub>2</sub>SO<sub>4</sub></td>
</tr>
</table>

<p>pH is adjusted to final value with NaOH (1 N)</p>

<hr align=left size=1 width="33%">

<p><b>Source:</b> Spanswick RM (1972) Evidence for an electrogenic ion pump in <i>Nitella translucens</i>. I. The effects of pH, K<sup>+</sup>, Na<sup>+</sup>,
light and temperature on the membrane potential and resistence. Biochimica
Biophysica Acta 288:74-89</p>

File:Nitella ligature.png

2013-04-01T11:11:15Z

Rlew:

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2013-04-01T11:10:48Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), MOPS buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
[[File:Nitella_ligature.png|300px|right]]
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming (ligature experiments by Varley in 1844 are shown in the watercolor [right]). What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://eol.org/pages/11583/overview)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie (starring Chara in the culture tank in your BPHS4090 teaching lab). The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

<h1> Appendix Three: APW (artificial pond water) Make-Up </h1>

<p>Make-Up of APW (artificial pond water)</p>

<p>Artificial pond water (APW)
is used to mimic the concentrations of major ions normally found in
freshwater environments, where algae typically grow. The APW stocks are stored
as X100 concentrated solutions in a refrigerator. When diluted to final
concentration, the pH is adjusted to the appropriate level with NaOH. APW 5 is
adjusted to pH 5.0, APW 6 is adjusted to pH 6.0, etc.</p>

<p>X100 stocks are made-up as 100 mM final:</p>

<table border=1 width=600>
<tr><td><b>Salts</b></td><td>Molecular Weight</td><td>grams/100 ml</td>
</tr>
<tr><td>Potassium chloride (KCl)</td><td>74.56</td><td>0.746</td>
</tr>
<tr><td>Calcium chloride, dihydrate (CaCl<sub>2</sub> 2H<sub>2</sub>O)</td><td>147.02</td><td>1.470</td>
</tr>
<tr><td>Magnesium chloride, hexahydrate (MgCl<sub>2</sub> 6H<sub>2</sub>O)</td><td>203.31</td><td>2.033</td>
</tr>
<tr><td>Sodium chloride (NaCl)</td><td>58.44</td><td>0.584</td>
</tr>
<tr><td>Sodium sulfate (Na<sub>2</sub>SO<sub>4</sub>) (for APW10)</td><td>142.04</td><td>1.420</td>
</tr>
<tr><td><b>Buffers</b></td><td></td><td></td>
</tr>
<tr><td>MES (pK<sub>a</sub> 6.15)</td><td>195.23</td><td>1.952</td>
</tr>
<tr><td>MOPS (pK<sub>a</sub> 7.20) </td><td>209.26</td><td>2.092</td>
</tr>
<tr><td>TES (pK<sub>a</sub> 7.50)</td><td>229.25</td><td>2.293</td>
</tr>
<tr><td>TAPS (pK<sub>a</sub> 8.40)</td><td>243.20</td><td>2.432</td>
</tr>
<tr><td>CAPS (pK<sub>a</sub> 10.40)</td><td>221.32</td><td>2.213</td>
</tr>
</table>

<p>For final solutions (final volume 1 liter)</p>

<table border=1 width=600>
<tr><td>100 mM Stock</td><td>KCl</td><td>CaCl<sub>2</sub></td><td>MgCl<sub>2</sub></td><td>NaCl</td><td>Buffer</td>
</tr>
<tr><td><p></p></td><td></td><td></td><td></td><td></td><td align=right>(in ml/1000 ml)</td>
</tr>
<tr><td>APW 5</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 6</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 7</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MOPS</td>
</tr>
<tr><td>APW 8</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TES</td>
</tr>
<tr><td>APW 9</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TAPS</td>
</tr>
<tr><td>APW 10</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 CAPS plus 0.05 Na<sub>2</sub>SO<sub>4</sub></td>
</tr>
</table>

<p>pH is adjusted to final value with NaOH (1 N)</p>

<hr align=left size=1 width="33%">

<p><b>Source:</b> Spanswick RM (1972) Evidence for an electrogenic ion pump in <i>Nitella translucens</i>. I. The effects of pH, K<sup>+</sup>, Na<sup>+</sup>,
light and temperature on the membrane potential and resistence. Biochimica
Biophysica Acta 288:74-89</p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-10-29T18:42:24Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), MOPS buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://eol.org/pages/11583/overview)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie (starring Chara in the culture tank in your BPHS4090 teaching lab). The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

<h1> Appendix Three: APW (artificial pond water) Make-Up </h1>

<p>Make-Up of APW (artificial pond water)</p>

<p>Artificial pond water (APW)
is used to mimic the concentrations of major ions normally found in
freshwater environments, where algae typically grow. The APW stocks are stored
as X100 concentrated solutions in a refrigerator. When diluted to final
concentration, the pH is adjusted to the appropriate level with NaOH. APW 5 is
adjusted to pH 5.0, APW 6 is adjusted to pH 6.0, etc.</p>

<p>X100 stocks are made-up as 100 mM final:</p>

<table border=1 width=600>
<tr><td><b>Salts</b></td><td>Molecular Weight</td><td>grams/100 ml</td>
</tr>
<tr><td>Potassium chloride (KCl)</td><td>74.56</td><td>0.746</td>
</tr>
<tr><td>Calcium chloride, dihydrate (CaCl<sub>2</sub> 2H<sub>2</sub>O)</td><td>147.02</td><td>1.470</td>
</tr>
<tr><td>Magnesium chloride, hexahydrate (MgCl<sub>2</sub> 6H<sub>2</sub>O)</td><td>203.31</td><td>2.033</td>
</tr>
<tr><td>Sodium chloride (NaCl)</td><td>58.44</td><td>0.584</td>
</tr>
<tr><td>Sodium sulfate (Na<sub>2</sub>SO<sub>4</sub>) (for APW10)</td><td>142.04</td><td>1.420</td>
</tr>
<tr><td><b>Buffers</b></td><td></td><td></td>
</tr>
<tr><td>MES (pK<sub>a</sub> 6.15)</td><td>195.23</td><td>1.952</td>
</tr>
<tr><td>MOPS (pK<sub>a</sub> 7.20) </td><td>209.26</td><td>2.092</td>
</tr>
<tr><td>TES (pK<sub>a</sub> 7.50)</td><td>229.25</td><td>2.293</td>
</tr>
<tr><td>TAPS (pK<sub>a</sub> 8.40)</td><td>243.20</td><td>2.432</td>
</tr>
<tr><td>CAPS (pK<sub>a</sub> 10.40)</td><td>221.32</td><td>2.213</td>
</tr>
</table>

<p>For final solutions (final volume 1 liter)</p>

<table border=1 width=600>
<tr><td>100 mM Stock</td><td>KCl</td><td>CaCl<sub>2</sub></td><td>MgCl<sub>2</sub></td><td>NaCl</td><td>Buffer</td>
</tr>
<tr><td><p></p></td><td></td><td></td><td></td><td></td><td align=right>(in ml/1000 ml)</td>
</tr>
<tr><td>APW 5</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 6</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 7</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MOPS</td>
</tr>
<tr><td>APW 8</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TES</td>
</tr>
<tr><td>APW 9</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TAPS</td>
</tr>
<tr><td>APW 10</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 CAPS plus 0.05 Na<sub>2</sub>SO<sub>4</sub></td>
</tr>
</table>

<p>pH is adjusted to final value with NaOH (1 N)</p>

<hr align=left size=1 width="33%">

<p><b>Source:</b> Spanswick RM (1972) Evidence for an electrogenic ion pump in <i>Nitella translucens</i>. I. The effects of pH, K<sup>+</sup>, Na<sup>+</sup>,
light and temperature on the membrane potential and resistence. Biochimica
Biophysica Acta 288:74-89</p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-10-29T13:56:09Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://eol.org/pages/11583/overview)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie (starring Chara in the culture tank in your BPHS4090 teaching lab). The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

<h1> Appendix Three: APW (artificial pond water) Make-Up </h1>

<p>Make-Up of APW (artificial pond water)</p>

<p>Artificial pond water (APW)
is used to mimic the concentrations of major ions normally found in
freshwater environments, where algae typically grow. The APW stocks are stored
as X100 concentrated solutions in a refrigerator. When diluted to final
concentration, the pH is adjusted to the appropriate level with NaOH. APW 5 is
adjusted to pH 5.0, APW 6 is adjusted to pH 6.0, etc.</p>

<p>X100 stocks are made-up as 100 mM final:</p>

<table border=1 width=600>
<tr><td><b>Salts</b></td><td>Molecular Weight</td><td>grams/100 ml</td>
</tr>
<tr><td>Potassium chloride (KCl)</td><td>74.56</td><td>0.746</td>
</tr>
<tr><td>Calcium chloride, dihydrate (CaCl<sub>2</sub> 2H<sub>2</sub>O)</td><td>147.02</td><td>1.470</td>
</tr>
<tr><td>Magnesium chloride, hexahydrate (MgCl<sub>2</sub> 6H<sub>2</sub>O)</td><td>203.31</td><td>2.033</td>
</tr>
<tr><td>Sodium chloride (NaCl)</td><td>58.44</td><td>0.584</td>
</tr>
<tr><td>Sodium sulfate (Na<sub>2</sub>SO<sub>4</sub>) (for APW10)</td><td>142.04</td><td>1.420</td>
</tr>
<tr><td><b>Buffers</b></td><td></td><td></td>
</tr>
<tr><td>MES (pK<sub>a</sub> 6.15)</td><td>195.23</td><td>1.952</td>
</tr>
<tr><td>MOPS (pK<sub>a</sub> 7.20) </td><td>209.26</td><td>2.092</td>
</tr>
<tr><td>TES (pK<sub>a</sub> 7.50)</td><td>229.25</td><td>2.293</td>
</tr>
<tr><td>TAPS (pK<sub>a</sub> 8.40)</td><td>243.20</td><td>2.432</td>
</tr>
<tr><td>CAPS (pK<sub>a</sub> 10.40)</td><td>221.32</td><td>2.213</td>
</tr>
</table>

<p>For final solutions (final volume 1 liter)</p>

<table border=1 width=600>
<tr><td>100 mM Stock</td><td>KCl</td><td>CaCl<sub>2</sub></td><td>MgCl<sub>2</sub></td><td>NaCl</td><td>Buffer</td>
</tr>
<tr><td><p></p></td><td></td><td></td><td></td><td></td><td align=right>(in ml/1000 ml)</td>
</tr>
<tr><td>APW 5</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 6</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 7</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MOPS</td>
</tr>
<tr><td>APW 8</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TES</td>
</tr>
<tr><td>APW 9</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TAPS</td>
</tr>
<tr><td>APW 10</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 CAPS plus 0.05 Na<sub>2</sub>SO<sub>4</sub></td>
</tr>
</table>

<p>pH is adjusted to final value with NaOH (1 N)</p>

<hr align=left size=1 width="33%">

<p><b>Source:</b> Spanswick RM (1972) Evidence for an electrogenic ion pump in <i>Nitella translucens</i>. I. The effects of pH, K<sup>+</sup>, Na<sup>+</sup>,
light and temperature on the membrane potential and resistence. Biochimica
Biophysica Acta 288:74-89</p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-10-29T13:55:36Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://eol.org/pages/11583/overview)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie (starring Chara in the culture tank in your BPHS4090 teaching lab). The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

<h1> Appendix Three: APW (artificial pond water) Make-Up </h1>

<p>Make-Up of APW (artificial pond water)</p>

<p>Artificial pond water (APW)
is used to mimic the concentrations of major ions normally found in
freshwater environments, where algae typically grow. The APW stocks are stored
as X100 concentrated solutions in a refrigerator. When diluted to final
concentration, the pH is adjusted to the appropriate level with NaOH. APW 5 is
adjusted to pH 5.0, APW 6 is adjusted to pH 6.0, etc.</p>

<p>X100 stocks are made-up as 100 mM final:</p>

<table border=1 width=600>
<tr><td><b>Salts</b></td><td>Molecular Weight</td><td>grams/100 ml</td>
</tr>
<tr><td>Potassium chloride (KCl)</td><td>74.56</td><td>0.746</td>
</tr>
<tr><td>Calcium chloride, dihydrate (CaCl<sub>2</sub> 2H<sub>2</sub>O)</td><td>147.02</td><td>1.470</td>
</tr>
<tr><td>Magnesium chloride, hexahydrate (MgCl<sub>2</sub> 6H<sub>2</sub>O)</td><td>203.31</td><td>2.033</td>
</tr>
<tr><td>Sodium chloride (NaCl)</td><td>58.44</td><td>0.584</td>
</tr>
<tr><td>Sodium sulfate (Na<sub>2</sub>SO<sub>4</sub>) (for APW10)</td><td>142.04</td><td>1.420</td>
</tr>
<tr><td><b>Buffers</b></td><td></td><td></td>
</tr>
<tr><td>MES (pK<sub>a</sub> 6.15)</td><td>195.23</td><td>1.952</td>
</tr>
<tr><td>MOPS (pK<sub>a</sub> 7.20) </td><td>209.26</td><td>2.092</td>
</tr>
<tr><td>TES (pK<sub>a</sub> 7.50)</td><td>229.25</td><td>2.293</td>
</tr>
<tr><td>TAPS (pK<sub>a</sub> 8.40)</td><td>243.20</td><td>2.432</td>
</tr>
<tr><td>CAPS (pK<sub>a</sub> 10.40)</td><td>221.32</td><td>2.213</td>
</tr>
</table>

<p>For final solutions (final volume 1 liter)</p>

<table border=1 width=600>
<tr><td>100 mM Stock</td><td>KCl</td><td>CaCl<sub>2</sub></td><td>MgCl<sub>2</sub></td><td>NaCl</td><td>Buffer</td>
</tr>
<tr><td><p></p></td><td></td><td></td><td></td><td></td><td>(in ml/1000 ml)</td>
</tr>
<tr><td>APW 5</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 6</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 7</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MOPS</td>
</tr>
<tr><td>APW 8</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TES</td>
</tr>
<tr><td>APW 9</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TAPS</td>
</tr>
<tr><td>APW 10</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 CAPS plus 0.05 Na<sub>2</sub>SO<sub>4</sub></td>
</tr>
</table>

<p>pH is adjusted to final value with NaOH (1 N)</p>

<hr align=left size=1 width="33%">

<p><b>Source:</b> Spanswick RM (1972) Evidence for an electrogenic ion pump in <i>Nitella translucens</i>. I. The effects of pH, K<sup>+</sup>, Na<sup>+</sup>,
light and temperature on the membrane potential and resistence. Biochimica
Biophysica Acta 288:74-89</p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-10-29T13:55:10Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://eol.org/pages/11583/overview)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie (starring Chara in the culture tank in your BPHS4090 teaching lab). The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

<h1> Appendix Three: APW (artificial pond water) Make-Up </h1>

<p>Make-Up of APW (artificial pond water)</p>

<p>Artificial pond water (APW)
is used to mimic the concentrations of major ions normally found in
freshwater environments, where algae typically grow. The APW stocks are stored
as X100 concentrated solutions in a refrigerator. When diluted to final
concentration, the pH is adjusted to the appropriate level with NaOH. APW 5 is
adjusted to pH 5.0, APW 6 is adjusted to pH 6.0, etc.</p>

<p>X100 stocks are made-up as 100 mM final:</p>

<table border=1 width=600>
<tr><td><b>Salts</b></td><td>Molecular Weight</td><td>grams/100 ml</td>
</tr>
<tr><td>Potassium chloride (KCl)</td><td>74.56</td><td>0.746</td>
</tr>
<tr><td>Calcium chloride, dihydrate (CaCl<sub>2</sub> 2H<sub>2</sub>O)</td><td>147.02</td><td>1.470</td>
</tr>
<tr><td>Magnesium chloride, hexahydrate (MgCl<sub>2</sub> 6H<sub>2</sub>O)</td><td>203.31</td><td>2.033</td>
</tr>
<tr><td>Sodium chloride (NaCl)</td><td>58.44</td><td>0.584</td>
</tr>
<tr><td>Sodium sulfate (Na<sub>2</sub>SO<sub>4</sub>) (for APW10)</td><td>142.04</td><td>1.420</td>
</tr>
<tr><td><b>Buffers</b></td><td></td><td></td>
</tr>
<tr><td>MES (pK<sub>a</sub> 6.15)</td><td>195.23</td><td>1.952</td>
</tr>
<tr><td>MOPS (pK<sub>a</sub> 7.20) </td><td>209.26</td><td>2.092</td>
</tr>
<tr><td>TES (pK<sub>a</sub> 7.50)</td><td>229.25</td><td>2.293</td>
</tr>
<tr><td>TAPS (pK<sub>a</sub> 8.40)</td><td>243.20</td><td>2.432</td>
</tr>
<tr><td>CAPS (pK<sub>a</sub> 10.40)</td><td>221.32</td><td>2.213</td>
</tr>
</table>

<p>For final solutions (final volume 1 liter)</p>

<table border=1 width=600>
<tr><td>100 mM Stock</td><td>KCl</td><td>CaCl<sub>2</sub></td><td>MgCl<sub>2</sub></td><td>NaCl</td><td>Buffer</td>
</tr>
<tr><td><p></p></td><td></td><td></td><td>(in ml/1000 ml)</td>
</tr>
<tr><td>APW 5</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 6</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 7</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MOPS</td>
</tr>
<tr><td>APW 8</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TES</td>
</tr>
<tr><td>APW 9</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TAPS</td>
</tr>
<tr><td>APW 10</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 CAPS plus 0.05 Na<sub>2</sub>SO<sub>4</sub></td>
</tr>
</table>

<p>pH is adjusted to final value with NaOH (1 N)</p>

<hr align=left size=1 width="33%">

<p><b>Source:</b> Spanswick RM (1972) Evidence for an electrogenic ion pump in <i>Nitella translucens</i>. I. The effects of pH, K<sup>+</sup>, Na<sup>+</sup>,
light and temperature on the membrane potential and resistence. Biochimica
Biophysica Acta 288:74-89</p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-10-26T16:10:07Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://eol.org/pages/11583/overview)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie (starring Chara in the culture tank in your BPHS4090 teaching lab). The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

<h1> Appendix Three: APW (artificial pond water) Make-Up </h1>

<p>Make-Up of APW (artificial pond water)</p>

<p>Artificial pond water (APW)
is used to mimic the concentrations of major ions normally found in
freshwater environments, where algae typically grow. The APW stocks are stored
as X100 concentrated solutions in a refrigerator. When diluted to final
concentration, the pH is adjusted to the appropriate level with NaOH. APW 5 is
adjusted to pH 5.0, APW 6 is adjusted to pH 6.0, etc.</p>

<p>X100 stocks are made-up as 100 mM final:</p>

<table border=1 width=600>
<tr><td><b>Salts</b></td><td>Molecular Weight</td><td>grams/100 ml</td>
</tr>
<tr><td>Potassium chloride (KCl)</td><td>74.56</td><td>0.746</td>
</tr>
<tr><td>Calcium chloride, dihydrate (CaCl<sub>2</sub> 2H<sub>2</sub>O)</td><td>147.02</td><td>1.470</td>
</tr>
<tr><td>Magnesium chloride, hexahydrate (MgCl<sub>2</sub> 6H<sub>2</sub>O)</td><td>203.31</td><td>2.033</td>
</tr>
<tr><td>Sodium chloride (NaCl)</td><td>58.44</td><td>0.584</td>
</tr>
<tr><td>Sodium sulfate (Na<sub>2</sub>SO<sub>4</sub>) (for APW10)</td><td>142.04</td><td>1.420</td>
</tr>
<tr><td><b>Buffers</b></td><td></td><td></td>
</tr>
<tr><td>MES (pK<sub>a</sub> 6.15)</td><td>195.23</td><td>1.952</td>
</tr>
<tr><td>MOPS (pK<sub>a</sub> 7.20) </td><td>209.26</td><td>2.092</td>
</tr>
<tr><td>TES (pK<sub>a</sub> 7.50)</td><td>229.25</td><td>2.293</td>
</tr>
<tr><td>TAPS (pK<sub>a</sub> 8.40)</td><td>243.20</td><td>2.432</td>
</tr>
<tr><td>CAPS (pK<sub>a</sub> 10.40)</td><td>221.32</td><td>2.213</td>
</tr>
</table>

<p>For final solutions (final volume 1 liter)</p>

<table border=1 width=600>
<tr><td>100 mM Stock</td><td>KCl</td><td>CaCl<sub>2</sub></td><td>MgCl<sub>2</sub></td><td>NaCl</td><td>Buffer</td>
</tr>
<tr><td><p></p></td><td>(in ml/1000 ml)</td>
</tr>
<tr><td>APW 5</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 6</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 7</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MOPS</td>
</tr>
<tr><td>APW 8</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TES</td>
</tr>
<tr><td>APW 9</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TAPS</td>
</tr>
<tr><td>APW 10</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 CAPS plus 0.05 Na<sub>2</sub>SO<sub>4</sub></td>
</tr>
</table>

<p>pH is adjusted to final value with NaOH (1 N)</p>

<hr align=left size=1 width="33%">

<p><b>Source:</b> Spanswick RM (1972) Evidence for an electrogenic ion pump in <i>Nitella translucens</i>. I. The effects of pH, K<sup>+</sup>, Na<sup>+</sup>,
light and temperature on the membrane potential and resistence. Biochimica
Biophysica Acta 288:74-89</p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-10-26T16:09:08Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://eol.org/pages/11583/overview)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie (starring Chara in the culture tank in your BPHS4090 teaching lab). The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

<h1> Appendix Three: APW (artificial pond water) Make-Up </h1>

<p>Make-Up of APW (artificial pond water)</p>

<p>Artificial pond water (APW)
is used to mimic the concentrations of major ions normally found in
freshwater environments, where algae typically grow. The APW stocks are stored
as X100 concentrated solutions in a refrigerator. When diluted to final
concentration, the pH is adjusted to the appropriate level with NaOH. APW 5 is
adjusted to pH 5.0, APW 6 is adjusted to pH 6.0, etc.</p>

<p>X100 stocks are made-up as 100 mM final:</p>

<table border=1 width=600>
<tr><td><b>Salts</b></td><td>Molecular Weight</td><td>grams/100 ml</td>
</tr>
<tr><td>Potassium chloride (KCl)</td><td>74.56</td><td>0.746</td>
</tr>
<tr><td>Calcium chloride, dihydrate (CaCl<sub>2</sub> 2H<sub>2</sub>O)</td><td>147.02</td><td>1.470</td>
</tr>
<tr><td>Magnesium chloride, hexahydrate (MgCl<sub>2</sub> 6H<sub>2</sub>O)</td><td>203.31</td><td>2.033</td>
</tr>
<tr><td>Sodium chloride (NaCl)</td><td>58.44</td><td>0.584</td>
</tr>
<tr><td>Sodium sulfate (Na<sub>2</sub>SO<sub>4</sub>) (for APW10)</td><td>142.04</td><td>1.420</td>
</tr>
<tr><td><b>Buffers</b></td><td></td><td></td>
</tr>
<tr><td>MES (pK<sub>a</sub> 6.15)</td><td>195.23</td><td>1.952</td>
</tr>
<tr><td>MOPS (pK<sub>a</sub> 7.20) </td><td>209.26</td><td>2.092</td>
</tr>
<tr><td>TES (pK<sub>a</sub> 7.50)</td><td>229.25</td><td>2.293</td>
</tr>
<tr><td>TAPS (pK<sub>a</sub> 8.40)</td><td>243.20</td><td>2.432</td>
</tr>
<tr><td>CAPS (pK<sub>a</sub> 10.40)</td><td>221.32</td><td>2.213</td>
</tr>
</table>

<p>For final solutions (final volume 1 liter)</p>

<table border=1 width=600>
<tr><td>100 mM Stock</td><td>KCl</td><td>CaCl<sub>2</sub></td><td>MgCl<sub>2</sub></td><td>NaCl</td><td>Buffer</td>
</tr><tr><td></td><td>(in ml/1000 ml)</td>
</tr>
<tr><td>APW 5</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 6</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 7</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MOPS</td>
</tr>
<tr><td>APW 8</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TES</td>
</tr>
<tr><td>APW 9</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TAPS</td>
</tr>
<tr><td>APW 10</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 CAPS plus 0.05 Na<sub>2</sub>SO<sub>4</sub></td>
</tr>
</table>

<p>pH is adjusted to final value with NaOH (1 N)</p>

<hr align=left size=1 width="33%">

<p><b>Source:</b> Spanswick RM (1972) Evidence for an electrogenic ion pump in <i>Nitella translucens</i>. I. The effects of pH, K<sup>+</sup>, Na<sup>+</sup>,
light and temperature on the membrane potential and resistence. Biochimica
Biophysica Acta 288:74-89</p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-10-26T16:01:25Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://eol.org/pages/11583/overview)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie (starring Chara in the culture tank in your BPHS4090 teaching lab). The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

<h1> Appendix Three: APW (artificial pond water) Make-Up </h1>

<p>Make-Up of APW (artificial pond water)</p>

<p>Artificial pond water (APW)
is used to mimic the concentrations of major ions normally found in
freshwater environments, where algae typically grow. The APW stocks are stored
as X100 concentrated solutions in a refrigerator. When diluted to final
concentration, the pH is adjusted to the appropriate level with NaOH. APW 5 is
adjusted to pH 5.0, APW 6 is adjusted to pH 6.0, etc.</p>

<p>X100 stocks are made-up as 100 mM final:</p>

<table border=1 width=600>
<tr><td><b>Salts</b></td><td>Molecular Weight</td><td>grams/100 ml</td>
</tr>
<tr><td>Potassium chloride (KCl)</td><td>74.56</td><td>0.746</td>
</tr>
<tr><td>Calcium chloride, dihydrate (CaCl<sub>2</sub> 2H<sub>2</sub>O)</td><td>147.02</td><td>1.470</td>
</tr>
<tr><td>Magnesium chloride, hexahydrate (MgCl<sub>2</sub> 6H<sub>2</sub>O)</td><td>203.31</td><td>2.033</td>
</tr>
<tr><td>Sodium chloride (NaCl)</td><td>58.44</td><td>0.584</td>
</tr>
<tr><td>Sodium sulfate (Na<sub>2</sub>SO<sub>4</sub>) (for APW10)</td><td>142.04</td><td>1.420</td>
</tr>
<tr><td><b>Buffers</b></td><td></td><td></td>
</tr>
<tr><td>MES (pK<sub>a</sub> 6.15)</td><td>195.23</td><td>1.952</td>
</tr>
<tr><td>MOPS (pK<sub>a</sub> 7.20) </td><td>209.26</td><td>2.092</td>
</tr>
<tr><td>TES (pK<sub>a</sub> 7.50)</td><td>229.25</td><td>2.293</td>
</tr>
<tr><td>TAPS (pK<sub>a</sub> 8.40)</td><td>243.20</td><td>2.432</td>
</tr>
<tr><td>CAPS (pK<sub>a</sub> 10.40)</td><td>221.32</td><td>2.213</td>
</tr>
</table>

<p>For final solutions (final volume 1 liter)</p>

<table border=1 width=600>
<tr><td>100 mM Stock</td><td>KCl</td><td>CaCl<sub>2</sub></td><td>MgCl<sub>2</sub></td><td>NaCl</td><td>Buffer</td>
</tr><tr><td></td><td><p>(in ml/1000 ml)</p></td>
</tr>
<tr><td>APW 5</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 6</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MES</td>
</tr>
<tr><td>APW 7</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 MOPS</td>
</tr>
<tr><td>APW 8</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TES</td>
</tr>
<tr><td>APW 9</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 TAPS</td>
</tr>
<tr><td>APW 10</td><td>0.1</td><td>0.1</td><td>0.1</td><td>0.5</td><td>1.0 CAPS plus 0.05 Na<sub>2</sub>SO<sub>4</sub></td>
</tr>
</table>

<p>pH is adjusted to final value with NaOH (1 N)</p>

<hr align=left size=1 width="33%">

<p><b>Source:</b> Spanswick RM (1972) Evidence for an electrogenic ion pump in <i>Nitella translucens</i>. I. The effects of pH, K<sup>+</sup>, Na<sup>+</sup>,
light and temperature on the membrane potential and resistence. Biochimica
Biophysica Acta 288:74-89</p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-10-26T16:00:59Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://eol.org/pages/11583/overview)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie (starring Chara in the culture tank in your BPHS4090 teaching lab). The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

<h1> Appendix Three: APW (artificial pond water) Make-Up </h1>

Main Page/BPHS 4090

2012-09-20T20:37:47Z

Rlew:

<table width=400 align=right><td>
[[File:Raman_logo2.png|350px|border|center]]

</td></table>

<h1>PHYS 4090 4.0 BioPhysics II</h1>
<br clear=right>
<h2>Course Director</h2>
<p>Dr. Christopher Bergevin</p>
<p>240 PSE</p>
<p>cberge@yorku.ca</p>
<p></p>
<p></p>

<h2>Laboratory Technologists </h2>
<table width=400>
<tr><td> Matthew George</td> <td> Nick Balaskas </td> </tr>
<tr><td> 122 PSE</td> <td> 122 PSE </td> </tr>
<tr><td> mgeorge@yorku.ca </td> <td> nickolaos@yorku.ca </td> </tr>
</table>

<h2>Prerequisite</h2>
<ul>
<li>BPHS 2090 2.0</li>
<li>PHYS 2020 3.0</li>
<li>PHYS 2060 3.0</li>
</ul>

<h1>Laboratory Manual</h1>
<ul>
<li> Sept. 10: [[Main Page/BPHS 4090/microscopy I|Transmitted Light Microscopy]] <b> </b> </li>
<li> Sept. 17: [[Main Page/BPHS 4090/microscopy II|Contrast Modes in Microscopy]] <b> </b> </li>
<li> Sept. 24: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 1: [[Main Page/BPHS 4090/Optical Tweezers of Onions|Optical Tweezers of Onion Cells]] <b></b> </li>
<li> Oct. 15: [[Main Page/BPHS 4090/Choloplast Translocation|Light Induced Chloroplast Translocation]] <b> </b> </li>
<li> Oct. 22: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 29: [[Main Page/BPHS 4090/In-Vivo Spectrocopy|In-Vivo Spectroscopy]]<b></b></li>

<li> Nov. 5: [[Main Page/BPHS 4090/ElectroPhysiology of Chara revised|The Electrical Properties of ''Chara'']]<b> </b> </li>
<li> Nov. 12: [[Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy|Mapping a binding site using NMR spectroscopy]]<b> </b> </li>
<li> Nov. 19: Otoacoustic Emissions<b> </b> </li>
<li> Nov. 26: Student Project Presentations<b> </b> </li>
</ul>

Main Page/BPHS 4090

2012-09-20T20:36:58Z

Rlew:

<table width=400 align=right><td>
[[File:Raman_logo2.png|350px|border|center]]

</td></table>

<h1>PHYS 4090 4.0 BioPhysics II</h1>
<br clear=right>
<h2>Course Director</h2>
<p>Dr. Christopher Bergevin</p>
<p>240 PSE</p>
<p>cberge@yorku.ca</p>
<p></p>
<p></p>

<h2>Laboratory Technologists </h2>
<table width=400>
<tr><td> Matthew George</td> <td> Nick Balaskas </td> </tr>
<tr><td> 122 PSE</td> <td> 122PSE </td> </tr>
<tr><td> mgeorge@yorku.ca </td> <td> nickolaos@yorku.ca </td> </tr>
</table>

<h2>Prerequisite</h2>
<ul>
<li>BPHS 2090 2.0</li>
<li>PHYS 2020 3.0</li>
<li>PHYS 2060 3.0</li>
</ul>

<h1>Laboratory Manual</h1>
<ul>
<li> Sept. 10: [[Main Page/BPHS 4090/microscopy I|Transmitted Light Microscopy]] <b> </b> </li>
<li> Sept. 17: [[Main Page/BPHS 4090/microscopy II|Contrast Modes in Microscopy]] <b> </b> </li>
<li> Sept. 24: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 1: [[Main Page/BPHS 4090/Optical Tweezers of Onions|Optical Tweezers of Onion Cells]] <b></b> </li>
<li> Oct. 15: [[Main Page/BPHS 4090/Choloplast Translocation|Light Induced Chloroplast Translocation]] <b> </b> </li>
<li> Oct. 22: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 29: [[Main Page/BPHS 4090/In-Vivo Spectrocopy|In-Vivo Spectroscopy]]<b></b></li>

<li> Nov. 5: [[Main Page/BPHS 4090/ElectroPhysiology of Chara revised|The Electrical Properties of ''Chara'']]<b> </b> </li>
<li> Nov. 12: [[Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy|Mapping a binding site using NMR spectroscopy]]<b> </b> </li>
<li> Nov. 19: Otoacoustic Emissions<b> </b> </li>
<li> Nov. 26: Student Project Presentations<b> </b> </li>
</ul>

Main Page/BPHS 4090

2012-09-20T20:36:22Z

Rlew:

<table width=400 align=right><td>
[[File:Raman_logo2.png|350px|border|center]]

</td></table>
<br clear=right>
<h1>PHYS 4090 4.0 BioPhysics II</h1>

<h2>Course Director</h2>
<p>Dr. Christopher Bergevin</p>
<p>240 PSE</p>
<p>cberge@yorku.ca</p>
<p></p>
<p></p>

<h2>Laboratory Technologists </h2>
<table width=400>
<tr><td> Matthew George</td> <td> Nick Balaskas </td> </tr>
<tr><td> 122 PSE</td> <td> 122PSE </td> </tr>
<tr><td> mgeorge@yorku.ca </td> <td> nickolaos@yorku.ca </td> </tr>
</table>

<h2>Prerequisite</h2>
<ul>
<li>BPHS 2090 2.0</li>
<li>PHYS 2020 3.0</li>
<li>PHYS 2060 3.0</li>
</ul>

<h1>Laboratory Manual</h1>
<ul>
<li> Sept. 10: [[Main Page/BPHS 4090/microscopy I|Transmitted Light Microscopy]] <b> </b> </li>
<li> Sept. 17: [[Main Page/BPHS 4090/microscopy II|Contrast Modes in Microscopy]] <b> </b> </li>
<li> Sept. 24: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 1: [[Main Page/BPHS 4090/Optical Tweezers of Onions|Optical Tweezers of Onion Cells]] <b></b> </li>
<li> Oct. 15: [[Main Page/BPHS 4090/Choloplast Translocation|Light Induced Chloroplast Translocation]] <b> </b> </li>
<li> Oct. 22: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 29: [[Main Page/BPHS 4090/In-Vivo Spectrocopy|In-Vivo Spectroscopy]]<b></b></li>

<li> Nov. 5: [[Main Page/BPHS 4090/ElectroPhysiology of Chara revised|The Electrical Properties of ''Chara'']]<b> </b> </li>
<li> Nov. 12: [[Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy|Mapping a binding site using NMR spectroscopy]]<b> </b> </li>
<li> Nov. 19: Otoacoustic Emissions<b> </b> </li>
<li> Nov. 26: Student Project Presentations<b> </b> </li>
</ul>

Main Page/BPHS 4090

2012-09-20T20:35:34Z

Rlew:

<table width=400 align=right><td>
[[File:Raman_logo2.png|350px|border|center]]

</td></table>

<h1>PHYS 4090 4.0 BioPhysics II</h1>
<br clear=right>
<h2>Course Director</h2>
<p>Dr. Christopher Bergevin</p>
<p>240 PSE</p>
<p>cberge@yorku.ca</p>
<p></p>
<p></p>

<h2>Laboratory Technologists </h2>
<table width=400>
<tr><td> Matthew George</td> <td> Nick Balaskas </td> </tr>
<tr><td> 122 PSE</td> <td> 122PSE </td> </tr>
<tr><td> mgeorge@yorku.ca </td> <td> nickolaos@yorku.ca </td> </tr>
</table>

<h2>Prerequisite</h2>
<ul>
<li>BPHS 2090 2.0</li>
<li>PHYS 2020 3.0</li>
<li>PHYS 2060 3.0</li>
</ul>

<h1>Laboratory Manual</h1>
<ul>
<li> Sept. 10: [[Main Page/BPHS 4090/microscopy I|Transmitted Light Microscopy]] <b> </b> </li>
<li> Sept. 17: [[Main Page/BPHS 4090/microscopy II|Contrast Modes in Microscopy]] <b> </b> </li>
<li> Sept. 24: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 1: [[Main Page/BPHS 4090/Optical Tweezers of Onions|Optical Tweezers of Onion Cells]] <b></b> </li>
<li> Oct. 15: [[Main Page/BPHS 4090/Choloplast Translocation|Light Induced Chloroplast Translocation]] <b> </b> </li>
<li> Oct. 22: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 29: [[Main Page/BPHS 4090/In-Vivo Spectrocopy|In-Vivo Spectroscopy]]<b></b></li>

<li> Nov. 5: [[Main Page/BPHS 4090/ElectroPhysiology of Chara revised|The Electrical Properties of ''Chara'']]<b> </b> </li>
<li> Nov. 12: [[Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy|Mapping a binding site using NMR spectroscopy]]<b> </b> </li>
<li> Nov. 19: Otoacoustic Emissions<b> </b> </li>
<li> Nov. 26: Student Project Presentations<b> </b> </li>
</ul>

Main Page/BPHS 4090

2012-09-20T20:35:23Z

Rlew:

<table width=400 align=right><td>
[[File:Raman_logo2.png|300px|border|center]]

</td></table>

<h1>PHYS 4090 4.0 BioPhysics II</h1>
<br clear=right>
<h2>Course Director</h2>
<p>Dr. Christopher Bergevin</p>
<p>240 PSE</p>
<p>cberge@yorku.ca</p>
<p></p>
<p></p>

<h2>Laboratory Technologists </h2>
<table width=400>
<tr><td> Matthew George</td> <td> Nick Balaskas </td> </tr>
<tr><td> 122 PSE</td> <td> 122PSE </td> </tr>
<tr><td> mgeorge@yorku.ca </td> <td> nickolaos@yorku.ca </td> </tr>
</table>

<h2>Prerequisite</h2>
<ul>
<li>BPHS 2090 2.0</li>
<li>PHYS 2020 3.0</li>
<li>PHYS 2060 3.0</li>
</ul>

<h1>Laboratory Manual</h1>
<ul>
<li> Sept. 10: [[Main Page/BPHS 4090/microscopy I|Transmitted Light Microscopy]] <b> </b> </li>
<li> Sept. 17: [[Main Page/BPHS 4090/microscopy II|Contrast Modes in Microscopy]] <b> </b> </li>
<li> Sept. 24: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 1: [[Main Page/BPHS 4090/Optical Tweezers of Onions|Optical Tweezers of Onion Cells]] <b></b> </li>
<li> Oct. 15: [[Main Page/BPHS 4090/Choloplast Translocation|Light Induced Chloroplast Translocation]] <b> </b> </li>
<li> Oct. 22: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 29: [[Main Page/BPHS 4090/In-Vivo Spectrocopy|In-Vivo Spectroscopy]]<b></b></li>

<li> Nov. 5: [[Main Page/BPHS 4090/ElectroPhysiology of Chara revised|The Electrical Properties of ''Chara'']]<b> </b> </li>
<li> Nov. 12: [[Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy|Mapping a binding site using NMR spectroscopy]]<b> </b> </li>
<li> Nov. 19: Otoacoustic Emissions<b> </b> </li>
<li> Nov. 26: Student Project Presentations<b> </b> </li>
</ul>

Main Page/BPHS 4090

2012-09-20T20:35:10Z

Rlew:

<table width=400 align=right><td>
[[File:Raman_logo2.png|400px|border|center]]

</td></table>

<h1>PHYS 4090 4.0 BioPhysics II</h1>
<br clear=right>
<h2>Course Director</h2>
<p>Dr. Christopher Bergevin</p>
<p>240 PSE</p>
<p>cberge@yorku.ca</p>
<p></p>
<p></p>

<h2>Laboratory Technologists </h2>
<table width=400>
<tr><td> Matthew George</td> <td> Nick Balaskas </td> </tr>
<tr><td> 122 PSE</td> <td> 122PSE </td> </tr>
<tr><td> mgeorge@yorku.ca </td> <td> nickolaos@yorku.ca </td> </tr>
</table>

<h2>Prerequisite</h2>
<ul>
<li>BPHS 2090 2.0</li>
<li>PHYS 2020 3.0</li>
<li>PHYS 2060 3.0</li>
</ul>

<h1>Laboratory Manual</h1>
<ul>
<li> Sept. 10: [[Main Page/BPHS 4090/microscopy I|Transmitted Light Microscopy]] <b> </b> </li>
<li> Sept. 17: [[Main Page/BPHS 4090/microscopy II|Contrast Modes in Microscopy]] <b> </b> </li>
<li> Sept. 24: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 1: [[Main Page/BPHS 4090/Optical Tweezers of Onions|Optical Tweezers of Onion Cells]] <b></b> </li>
<li> Oct. 15: [[Main Page/BPHS 4090/Choloplast Translocation|Light Induced Chloroplast Translocation]] <b> </b> </li>
<li> Oct. 22: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 29: [[Main Page/BPHS 4090/In-Vivo Spectrocopy|In-Vivo Spectroscopy]]<b></b></li>

<li> Nov. 5: [[Main Page/BPHS 4090/ElectroPhysiology of Chara revised|The Electrical Properties of ''Chara'']]<b> </b> </li>
<li> Nov. 12: [[Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy|Mapping a binding site using NMR spectroscopy]]<b> </b> </li>
<li> Nov. 19: Otoacoustic Emissions<b> </b> </li>
<li> Nov. 26: Student Project Presentations<b> </b> </li>
</ul>

Main Page/BPHS 4090

2012-09-20T20:34:52Z

Rlew:

<table width=400 align=right><td>
[[File:Raman_logo2.png|500px|border|center]]

</td></table>

<h1>PHYS 4090 4.0 BioPhysics II</h1>
<br clear=right>
<h2>Course Director</h2>
<p>Dr. Christopher Bergevin</p>
<p>240 PSE</p>
<p>cberge@yorku.ca</p>
<p></p>
<p></p>

<h2>Laboratory Technologists </h2>
<table width=400>
<tr><td> Matthew George</td> <td> Nick Balaskas </td> </tr>
<tr><td> 122 PSE</td> <td> 122PSE </td> </tr>
<tr><td> mgeorge@yorku.ca </td> <td> nickolaos@yorku.ca </td> </tr>
</table>

<h2>Prerequisite</h2>
<ul>
<li>BPHS 2090 2.0</li>
<li>PHYS 2020 3.0</li>
<li>PHYS 2060 3.0</li>
</ul>

<h1>Laboratory Manual</h1>
<ul>
<li> Sept. 10: [[Main Page/BPHS 4090/microscopy I|Transmitted Light Microscopy]] <b> </b> </li>
<li> Sept. 17: [[Main Page/BPHS 4090/microscopy II|Contrast Modes in Microscopy]] <b> </b> </li>
<li> Sept. 24: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 1: [[Main Page/BPHS 4090/Optical Tweezers of Onions|Optical Tweezers of Onion Cells]] <b></b> </li>
<li> Oct. 15: [[Main Page/BPHS 4090/Choloplast Translocation|Light Induced Chloroplast Translocation]] <b> </b> </li>
<li> Oct. 22: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 29: [[Main Page/BPHS 4090/In-Vivo Spectrocopy|In-Vivo Spectroscopy]]<b></b></li>

<li> Nov. 5: [[Main Page/BPHS 4090/ElectroPhysiology of Chara revised|The Electrical Properties of ''Chara'']]<b> </b> </li>
<li> Nov. 12: [[Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy|Mapping a binding site using NMR spectroscopy]]<b> </b> </li>
<li> Nov. 19: Otoacoustic Emissions<b> </b> </li>
<li> Nov. 26: Student Project Presentations<b> </b> </li>
</ul>

Main Page/BPHS 4090

2012-09-20T20:32:47Z

Rlew:

<table width=400 align=center><td>
<p align=justify>[[File:Raman_logo2.png|500px|border|center]]
<br clear=right>
</p>
</td></table>

<h1>PHYS 4090 4.0 BioPhysics II</h1>

<h2>Course Director</h2>
<p>Dr. Christopher Bergevin</p>
<p>240 PSE</p>
<p>cberge@yorku.ca</p>
<p></p>
<p></p>

<h2>Laboratory Technologists </h2>
<table width=400>
<tr><td> Matthew George</td> <td> Nick Balaskas </td> </tr>
<tr><td> 122 PSE</td> <td> 122PSE </td> </tr>
<tr><td> mgeorge@yorku.ca </td> <td> nickolaos@yorku.ca </td> </tr>
</table>

<h2>Prerequisite</h2>
<ul>
<li>BPHS 2090 2.0</li>
<li>PHYS 2020 3.0</li>
<li>PHYS 2060 3.0</li>
</ul>

<h1>Laboratory Manual</h1>
<ul>
<li> Sept. 10: [[Main Page/BPHS 4090/microscopy I|Transmitted Light Microscopy]] <b> </b> </li>
<li> Sept. 17: [[Main Page/BPHS 4090/microscopy II|Contrast Modes in Microscopy]] <b> </b> </li>
<li> Sept. 24: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 1: [[Main Page/BPHS 4090/Optical Tweezers of Onions|Optical Tweezers of Onion Cells]] <b></b> </li>
<li> Oct. 15: [[Main Page/BPHS 4090/Choloplast Translocation|Light Induced Chloroplast Translocation]] <b> </b> </li>
<li> Oct. 22: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 29: [[Main Page/BPHS 4090/In-Vivo Spectrocopy|In-Vivo Spectroscopy]]<b></b></li>

<li> Nov. 5: [[Main Page/BPHS 4090/ElectroPhysiology of Chara revised|The Electrical Properties of ''Chara'']]<b> </b> </li>
<li> Nov. 12: [[Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy|Mapping a binding site using NMR spectroscopy]]<b> </b> </li>
<li> Nov. 19: Otoacoustic Emissions<b> </b> </li>
<li> Nov. 26: Student Project Presentations<b> </b> </li>
</ul>

File:Diatom Slide.png

2012-08-22T11:23:14Z

Rlew: A revised image showing the slide, the resolutions and a X40 DIC image of one of the diatoms to demonstrate resolution at 0.72 microns.

A revised image showing the slide, the resolutions and a X40 DIC image of one of the diatoms to demonstrate resolution at 0.72 microns.

Main Page/BPHS 4090/microscopy I

2012-08-22T11:21:19Z

Rlew:

<h1>Required Components</h1>
<ol>
<li>[[Media:Diatom_Slide.png|Diatom slide]]</li>
<li>[[Media:Micrometer_Slide.JPG|Stage micrometer slide]]</li>
<li>[[Media:M1_Baboon.JPG|Stained and unstained baboon eye slides]]</li>
<li>[[Media:Slide_Tools.JPG|Fresh onion]]</li>
<li>[[Media:Slide_Tools.JPG|Knife and tweezers]]</li>
<li>[[Media:Slide_Tools.JPG|Toothpick or popsicle sticks]]</li>
<li>[[Media:Slide_Tools.JPG|Blank slides and coverslips]]</li>
<li>[[Media:Slide_Tools.JPG|Syringe of vaseline with pipet tip end]]</li>
<li>[[Media:Slide_Tools.JPG|Distilled water]]</li>
<li>[[Media:Immersion_Oil.JPG|Immersion oil]]</li>
<li>[[Media:CCD.JPG|Nikon upright microscope and CCD camera]]</li>
<li>USB stick for transferring data and images for processing and report generation</li>
</ol>

<h1>Objectives</h1>
<p>Students will get a refresher on the optical light path of microscopes as well as working with
digital acquisition software and data processing. Building off of previous knowledge, the
students will explore light contrast modes for unstained samples like darkfield imaging and
phase contrast imaging. In the first part of this lab these modes will be compared with standard
brightfield illumination on stained and unstained specimens. In the second part darkfield
imaging will be used to measure the cytoplasmic streaming rate of spherosomes in living onion
cells.</p>

<h1>Introduction</h1>
<p>Imaging has become a vital tool for researchers in virtually all aspects of modern biophysics.
Recent advances in microscope technology as well as labelling techniques and gene and protein
manipulation methods have led to breakthroughs in our understanding of biological processes.
In order to take advantage of these methods you, the biophysicist, need to understand some of
the fundamental techniques and concepts in microscopy. You already have experience with
microscopes, and this lab will build on that knowledge and get you familiar with more advanced
methods as well as working with acquisition software.</p>
<p>The brightfield (transmitted light) imaging mode is something you have already been exposed
to. Brightfield imaging mimics the human optical system and is measured in terms of colour and
light intensity absorbed by a sample. The contrast method is based off of colour specific
absorption of light by the dyes added into the specimen. When working with certain samples,
particularly live specimens, the addition of exogenous contrast agents isn’t always feasible as
they can potentially interfere with the organism or cells under observation. In this lab we will
focus on some specialized microscope techniques to look at unstained samples in a way that lets
you observe their morphology without the addition of foreign compounds. The imaging modes
you will be using are:</p>
<ol>
<li><b>Brightfield:</b> where samples contain little natural contrast and dyes are added to impart
artificial colour</li>
<li><b>Darkfield:</b> where photons observed are mostly those that have been scattered by structures
within the samples</li>
<li><b>Phase Contrast:</b> where optics will be used to visualize changes in the optical path length of
samples</li>
</ol>

<p>A refresher on optical microscopy and imaging theory is included in Appendix 1. Please refer to
this section to review some of the fundamental concepts in microscopy. If you feel comfortable
with this content you do not have to read through it thoroughly. It has been included as
optional background material.</p>

<h2>Brightfield Staining</h2>
<p>Most biomedical microscopy specimens contain very little intrinsic contrast when viewed
directly with transmitted light. For example, to make specimens such as the tissue sections you
are using in this lab easier to visualize, stains and dyes are used. These compounds impart
additional contrast and enable discrimination of morphological features based on their
differential colour absorption across the visible spectrum.</p>
<p>By far, the most common type of stain used in biological microscopy is Haematoxylin & Eosin
(H&E). Haematoxylin stains basophilic chemical structures such as DNA in the nucleus dark
purple, while Eosin will stain the cytoplasm and extra-cellular regions a shade of pink. When
oxidized, haematoxylin forms a structure called haematein. Haematein is a compound that
forms strongly coloured complexes with metal ions, most notably Fe(III) and Al(III), and it is
these coloured complexes which give the blue/purple colouring seen in the nuclei of H&E
sections. H&E staining allows pathologists to observe the density and shape of nuclei in tissue
sections, and is the gold standard in the diagnosis of many types of Cancer.</p>
<p>Shown in Figure 1 is a sample of an H&E stained tissue section from a rabbit brain. A low
resolution image shows a brain tumour as the dark purple mass, and the zoom view shows the
border between tumour and healthy tissue. Note the dark purple staining of the cancerous
nuclei, and how densely packed they are.</p>

<table width=400 align=center><td>
<p align=justify>[[File:Fig1_stained.png|400px|border|center]]
<b>Figure 1 -</b> H&E stained section of mouse brain. Shown inset is a zoomed view showing the cluster of nuclei in a
brain tumour (dark purple patch in larger image).
<br clear=right>
</p>
</td></table>

<h2>Darkfield Microscopy</h2>
<p>Dark-field microscopy permits the detection of unstained small biological objects which
otherwise provide insufficient contrast under transmitted light observation. In a dark-field
microscope a special condenser or aperture stop is used to produce light rays that normally miss
being collected by the microscope objective’s collection NA. When this light interacts with a
scattering object such as a biological sample, light is scattered into the collection NA of the
objective and detected (Figure 2). While it is often the least used of the contrast modes you are
exploring, it is relatively simple to adapt a standard microscope for darkfield imaging, and the
images produced can be quite striking.</p>
<p>The aperture stop in a darkfield condenser causes light to focus on the specimen in a ‘hollow’
cone. The aperture stop is sized in such a way that this hollow cone is larger than the collection
NA of the objective being used, so most of the light put out by the lamp is not detected. Any
light that interacts with the sample will be scattered or refracted into the collection NA of the
objective being used (light grey region) and detected. Light that does not pass through the
sample does not enter the objective’s collection volume, and is thus not detected (gold region).</p>

<table width=600 align=center><td>
<p align=justify>[[File:Fig2_darkfield.png|400px|border|center]]
<b>Figure 2 -</b> Darkfield illumination path in a typical microscope. An aperture is used to create a ‘hollow’ beam of light
and is focused at the sample. Light that does not interact with the sample (scattering, refraction, reflection, etc) is
outside of the lens NA and is therefore not detected. Light that is scattered or refracted by the sample enters the
collection NA of the objective and is detected.
<br clear=right>
</p>
</td></table>
<p>Since there is very little scattering from the glass slides which the samples are mounted on, the image you see appears as a bright image on a dark background, hence the name 'darkfield' microscopy. This microscope method is very useful for viewing small bacteria as well as blood samples. You will also notice that if you are using slides that have not been thoroughly
cleaned the dust and dirt on the slide surface will produce a very strong darkfield signal. It is always good practice to clean microscope slides before observation, but this is even more important for darkfield imaging.</p>

<p>An ideal sample to demonstrate darkfield imaging is an onion cell, as it is readily available, cheap, and exhibits quite dynamic activity under darkfield illumination. The endoplasmic reticulum of onion cells forms a network of interconnected tubules within, within which highly refractile objects like mitochondria and sphereosomes are propelled by actin filaments within the cells cytoplasm. An excellent figure depicting the structure and location of this network in an onion cell is found in figure 1 of the paper
<i>"Onion Epidermal Cells in Relation to Microtubules, Microfilaments, and Intracellular Particle Movement”</i> by Stromgren-Allen and Brown, which should be provided to you as a separate pdf file (a copy is also stored in the ‘Onion Sample’ folder on the PC desktop). It is recommended that you read this paper as well as <i>“Isolation of Spherosomes (Oleosomes) from Onion, Cabbage,and Cottonseed Tissues”</i> by Yatsu et al, which includes an electron microscope and compositional analysis on the particles streaming around in this tubule network.</p>

<h2>Phase Contrast Microscopy </h2>
<p>Before getting into details regarding phase contrast imaging we should first review a concept
which you should have come across in your studies, the optical path length of an object. It is a
relatively simple concept, which essentially states that the optical path length of light traversing
an object is equal to the physical length travelled (L) multiplied by the index of refraction (n) of
the medium the light is travelling in. Thus, an object can be the same thickness throughout its
volume, but changes in the index of refraction make its optical path length vary. Our eyes are
sensitive to amplitude (intensity), but not sensitive to changes in light phase induced by optical
path length differences. These are the changes we are visualizing in phase contrast imaging.</p>
<p align=center><i> Optical Path Length = n X L</i></p>
<p>Phase contrast microscopy produces image intensities that vary as a function of specimen
optical path length, with very dense regions (those having large path lengths) appearing darker
than the background. Alternatively, specimen features that have relatively low thickness values,
or a refractive index less than the surrounding medium, are rendered much lighter when
superimposed on the standard (positive) phase contrast (grey) background. Remember; as light
travels through a medium other than vacuum, interaction with this medium causes its amplitude
and phase to change in a way which depends on properties of the medium (Figure 3). Changes
in amplitude give rise to absorption of light, which gives rise to colours. The human eye
measures only the energy of light arriving on the retina, so changes in phase are not easily
observed, yet often these changes in phase carry a large amount of information.</p>

<table width=500 align=center><td>
<p align=justify>[[File:Fig3_phase.png|300px|border|center]]
<b>Figure 3 -</b> Two parallel light waves beginning perfectly in phase. The top wave travels through a homogenous
optical path while the bottom wave passes through a region where the index of refraction increases (boxed area).
The longer optical path of the bottom wave causes the waves to drift out of phase.
<br clear=right>
</p>
</td></table>

<p>In a typical phase contrast microscope the phase variations introduced by the sample are preserved by the instrument, but this information is usually lost in the process which measures the light. In order to make phase variations observable it is necessary to
combine the light passing through the sample with a reference wave and take advantage of constructive/destructive interference to highlight changes in phase. The resulting interference between the reference wave and the wave interacting with the sample reveals the optical path structure of the sample as intensity changes in the image observed or captured by a detector. The difference between a phase contrast and brightfield optical path are the addition of two matched phase rings etched accurately onto glass plates within the condenser as well as the objective lens (Figure 4). You can see that the rings are inverse patterns, and they must be
aligned co-axially to maximize the phase contrast effect (see Appendix 2 for this procedure).</p>

<table width=500 align=center><td>
<p align=justify>[[File:Fig4_condenser.png|300px|border|center]]
<b>Figure 4 -</b> Location of the condenser annulus and phase plate in a PC light path.
<br clear=right>
</p>
</td></table>

<h1>Methods</h1>
<p>A schematic of the microscope you will be using in this lab is shown in Figure 5. You can use this
figure as a reference when performing the alignment and configuration of the microscope
during the lab. Of particular note is to make sure you have the correct condenser fitted on the
scope for this lab, which is the phase contrast condenser (NOT the DIC condenser). For aligning
Kohler illumination use the centering screws on the base of the condenser mount (attached to
the condenser focus block. For aligning the phase rings use the centering screws attached to the
underside of the phase contrast condenser.</p>

<table width=400 align=center><td>
<p align=justify>[[File:Fig5_microscope.png|400px|border|center]]
<b>Figure 5 -</b> Schematic of the Nikon microscope you will be using in this lab. Refer to this figure to familiarize
yourself with the components and their locations.
<br clear=right>
</p>
</td></table>

<p>Before performing any measurements or turning anything on, take a moment to study the
labelled schematic of the upright microscope you are using. Below are some tips on how to
start up the image acquisition software as well as for working with oil immersion objectives.</p>
<ol>
<li><h3>Initial microscope setup – Kohler illumination</h3>
<ol style="list-style-type:lower-latin">
<li>Familiarize yourself with the microscope and the diagram included in this lab. We will
not be using the fluorescence filter cubes or mercury light source for this lab (these will
be used in the fluorescence lab).</li>
<li>Turn on the power to the microscope transmitted light source (power switch is on the
base of the microscope).</li>
<li>Select the 10x objective and select the H&E mouse brain slide and place it on the
specimen holder. Focus on the specimen.</li>
<li>Close the field diaphragm down until you can see its effect in the oculars.</li>
<li>Focus the condenser until you get a sharp image of the field diaphragm. You may need
to gently push the focus block while turning the height adjustment knob as it is a tight at
certain points in its travel.</li>
<li>When the field diaphragm is in focus, you will see a red-blue flare surrounding it and as
you open/close the diaphragm you will clearly see a circle getting larger/smaller.</li>
<li>Now that the condenser height is positioned properly use the adjustment screws on the
condenser to center the image of the diaphragm in the field of view.</li>
<li>Open the field diaphragm so that it is slightly larger than the field of view you are
looking at.</li>
<li> You should now be set up for Kohler illumination which illuminates the sample with a
uniform field of light. An example showing poor contrast and colour reproduction
caused by non Kohler illumination is shown below for reference:
<table width=400 align=center><td>
<p align=justify>[[File:Fig6_aligned.png|400px|border|center]]
<br clear=right>
</p>
</td></table>
</li>
<li>If you change to a different objective you may need to re-adjust the diaphragm size or
condenser height for optimal performance, but in general you should be ok for the rest
of the lab.</li>
<li>When aligning the phase contrast mode the Kohler illumination could shift slightly along
X & Y, which should not matter much as long as the condenser height is ok and the
aperture is opened slightly larger than the field of view.</li>
</li>
</ol>
<li><h3>Working with immersion objectives</h3>
<ol style="list-style-type:lower-latin">
<li>When working with immersion objectives be very careful with how much oil you use and clean the objective thoroughly when finished.</li>
<li>Also be careful not to get any oil on the non-immersion objectives. If this does happen,
remove the objective from the microscope and clean it with the lens cleaning solution.
Remove one of the microscope's eyepieces and use it to examine the lens and ensure it
is clean. Use the lens to look at a specimen to confirm it is clean. If it is still dirty, repeat
the cleaning procedure.</li>
<li>Move the stage down and use the light from the condenser as a reference for where to
place the drop on the specimen. You don’t need much immersion oil here, if you use
too much it will be a waste and make it more difficult to clean the slide when you are
finished.</li>
<li> Carefully finish rotating the 100x or 40x objective in place and re-focus on the sample.
At this point you can freely move around the slide, just try and keep the X-Y motion of
the sample to a slow and steady pace or else you will lose oil quickly and have to add
more. If you make slow, controlled motions the oil will not spread out as much.</li>
<li>After acquiring your images, lower the stage and remove the slide. Carefully clean the
slide and lens using a kimwipe and the supplied cleaning fluid.</li>
<li>When you are finished with the lab thoroughly clean any immersion objectives used as
well as slides that are part of the permanent lab (ie, micrometer, diatom slide, stained
slides). This is a critical step, as over time oil that is not cleaned can degrade image
quality and even render the very expensive objectives on the microscope useless. The
objectives will unscrew from the turret, and using one of the eyepieces you can examine
the lens after cleaning it to ensure there is no residual oil (ask your lab instructor to
show you how to do this if you are unsure).</li>
</ol>
<li><h3>Getting ready for acquisition and digital image capture.</h3>
<ol style="list-style-type:lower-latin">
<li>Turn the CCD on by pressing the power switch located on the top of the camera.</li>
<li>Launch the μManager software which will allow you to record data and adjust the
camera gain. Upon startup select the default configuration file in the splash screen that
starts up.
<table width=400 align=center><td>
<p align=justify>[[File:Fig7_micromanager.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>After startup, the main control screen for μManager should appear. From here you can
change the exposure settings on the CCD camera to optimize the signal by typing in an
exposure time (the units are milliseconds).
<table width=400 align=center><td>
<p align=justify>[[File:Fig8_config.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Rotate the eyepiece mounting block to the left to allow light to pass through to the CCD
camera.</li>
<li>Pressing the ‘live’ button in μManager should open a new window that shows a live
image of the light striking the CCD camera.</li>
<li>Pressing the ‘snap’ button will take a single snapshot which you can then save for later
processing.</li>
<li>Pressing the ‘Multi-D Acq’ button opens the multi dimensional acquisition dialogue from
which you can configure a time lapse acquisition (see below).</li>
</ol>
<li><h3>Setting up a timed acquisition in μManager</h3>
<ol style="list-style-type:lower-latin">
<li>For some experiments you will want to set up a timed acquisition so that you can make
a movie and dynamic measurements of samples. To do this open the multi-dimensional
acquisition dialogue and refer to the figure below to set relevant acquisition
parameters.
<ol style="list-style-type:lower-roman">
<li>Number = number of frames you want to acquire</li>
<li>Interval = time between frames (for setting up time-lapse)</li>
<li>Save Images = directory to save your images to (files will be automatically
numbered and timestamped)
<table width=400 align=center><td>
<p align=justify>[[File:Fig9_acquisition.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li></ol>
<li>It is a good idea to include parameters such as the CCD exposure time, microscope
objective, time delay and CCD bin settings in the file name or folder.</li>
<li>Be sure to change directories and file names between movies to keep the organization
of the files neat.</li>
<li>Pressing the ‘Acquire’ button will initiate the acquisition protocol using the parameters
you specified and the CCD gain currently being used.</li>
<li>Double check the folder after acquisition is complete to ensure all the files were
properly saved.</li>
</ol>
</ol>

<h1>Experiments</h1>
<p>There are a number of small exercises to be performed in this lab. The first few are relatively
straightforward, and shouldn’t take a significant amount of time to finish. Getting the onion
peel sample prepared correctly may take a few tries, and finding the most active cells and
recording the image sequences will also eat up a fair bit of time. If you do complete everything
early, there are a number of additional slides which you can look at in the slide cases.</p>
<p>First, set the microscope up for Kohler illumination, and follow the directions in Appendix 2 for
setting up the phase contrast alignment. Once you are finished with that, start by taking some
images of the stage micrometer so you can spatially calibrate the images. This will later allow
you to make measurements of object sizes and calculate cytoplasmic streaming speeds in the
onion cells. Follow through the steps and tips below.</p>

<ol>
<li><h3>Calibration images of micrometer slide & setting CCD exposure</h3>
<ol style="list-style-type:lower-latin">
<li>Acquire and save images of the metric scale using the 10x, 20x, 40x (air and oil),
100x oil objectives.</li>
<li>Acquire the oil immersion images last, and ensure there is no oil on the slide when
using the dry objectives.</li>
<li>Please ensure the slide is cleaned thoroughly and placed back in its case after you
are finished with it.</li>
<li>Place the micrometer slide on the microscope and rotate the eyepiece turret
counter clockwise (to the left if facing the microscope) to allow the light to pass
through to the CCD camera.</li>
<li>Note the CCD camera has an additional 2.5x doubling lens in front of it. Thus a 10x
image to your eye is magnified up to 25x on the CCD. In other words; the field of
view you see with your eyes when using a 100x objective is what the camera sees
when using the 40x objective.</li>
<li>Press the ‘Live’ button and prepare to adjust the CCD exposure setting.</li>
<li>Adjust the CCD exposure setting so that you get as bright an image as possible without saturating the CCD pixels. You can tell when you have a good exposure setting by observing the histogram in the lower portion of the μManager window.</li>
<li>Click on ‘Full’ and you will remove any digital contrast adjustment from the live
display. Set the exposure time so that the histogram in the graph window is
properly exposed like the example below (depending on your sample you may not
see two separate peaks as shown).</li>
<table width=500 align=center><td>
<p align=justify>[[File:Fig10_screenshot.png|500px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>You can use the slider bars to set the black and white levels of the image (applying
contrast), but it is always best to start by displaying the full data range.</li>
<li>When you are happy with the exposure setting you can press the ‘Snap’ button to
take a single image which will open in a separate window. You can then save this
image for later processing in ImageJ.</li>
<li>Use the objective magnification in the file name to make it easier for you to
correlate the proper calibration images with the data you will need to calibrate. You
will calibrate the images after completing the experiments. Importing images
sequences and applying spatial calibrations in ImageJ is covered in Appendix 3.</li>
</ol>
<li><h3>Cheek epithelial cells in brightfield, darkfield and phase contrast</h3>
<ol style="list-style-type:lower-latin">
<li>By gently scraping a toothpick on the inside of your mouth you will remove a large
number of epithelial cells, which are ideal to demonstrate the difference between
the imaging modes being explored in this lab.</li>
<li>Transfer the material you scraped off to a glass slide.</li>
<li>Apply a thin bead of vaseline to the surface of a microscope slide, forming a square
approximately 1 cm2 in area around the cells.</li>
<li>Place a small drop of distilled water in the centre of the vaseline square.</li>
<li>Place a coverslip on top of it all and gently press down on it to seal the specimen.
This should prevent drying out of the cells during the experiment.</li>
<li>Observe the cells in darkfield, brightfield and phase contrast (Note; only the 10x
objective is equipped for phase contrast imaging).</li>
<li>Acquire images in each of these modes and compare them on the computer
monitor to see how the same structures appear visibly different depending on the
technique used.</li>
<li>Try to go for cells that are not folded and avoid cells that are clumped together. If
the cells are too tightly packed it is easy to prepare another one.</li>
<li>Most of the cells you will observe are dead or dying, which is why they come off so
easily. This makes them somewhat un-interesting for time-lapse imaging, but the
onion cells show significantly more activity.</li>
</ol>

<li><h3>Onion peel slide</h3>
<ol style="list-style-type:lower-latin">
<li>By cutting into an onion and carefully removing a thin transparent sheet of cells
from the interior layers we can catch some dynamic events.</li>
<li>After removing the onion peel, lay it flat on a microscope slide using tweezers and
toothpicks to gently pull it flat. It may help to put a small drop of water on the slide
first if you find that the peel is tearing easily because it sticks to the slide surface.</li>
<li>Cut the peel to approximately 1 cm<sup>2</sup> and apply vaseline around its border.</li>
<li>Place a drop of distilled water on the sample.</li>
<li>Seal with coverslip.</li>
<li>Observe in darkfield first using the 10x objective.</li>
<li>Fresh and healthy cells will have an intact cell wall, and clearly defined nucleus with
quite fast and dramatic streaming. Move around the sample looking for cells
exhibiting cytoplasmic streaming. You may need to switch to the 20x or 40x
objective to see the streaming better with your eyes.</li>
<li>If you don’t see any activity in this slide, make up another one from a different layer
in the onion. It may take several tries to get a fresh one. There is a sample movie in
the ‘Onion Sample’ folder on located on the computer desktop to demonstrate the
type of activity you are looking for. The movie was captured at 1 frame per second
and accurately shows the streaming speeds you should see when using the 10x and
20x objectives and the CCD camera. The 10x movie is what you would see using 25x
when looking with your eyes, and the 20x movie is equivalent to 50x when looking
with your eyes.</li>
<li>Try and find a cell away from any air bubbles or very bright objects.</li>
<li>Record movies of the cytoplasmic streaming using the 10x/Ph2 in transmitted light,
phase contrast and darkfield, as well as the 20x objectives which has no phase plate
and thus cannot image in phase contrast mode.</li>
<li>Open the ‘multi-dimensional acquisition’ window and select a folder to save your
data to. Be sure to indicate which objective you used in either the folder or file
name.</li>
<li>Acquiring 100 frames with a delay of 500 ms between frames should be more than
sufficient.</li>
</ol>

<li><h3>(OPTIONAL)Observe brightfield slides using all objectives (10x, 20x, 40x, 100x)</h3>
<ol style="list-style-type:lower-latin">
<li>Stained and unstained sections of a baboon eye should be provided to you by your
instructor.</li>
<li>You can clearly make out cellular structures, including the retinal layers opposite the
lens in the stained section. In the unstained section you will not see much other
than some natural pigments in the retinal layers.</li>
<li>You will also be provided two Pap smear test slides as an example of healthy and
diseased samples. Note the change in nuclear morphology between the samples.
Which sample looks ‘healthier’ to you?</li>
<li>A number of other slides are in the containers for you to explore and image if you
have time to come back to them. A list of what the specimens are and what stain
was used to prepare them is included in an Appendix 5.</li>
</ol>

<li><h3>(OPTIONAL) Using the diatom slide as well as the Pap smear slides compare the dry 40x objective with
the oil immersion 40x objective</h3>
<ol style="list-style-type:lower-latin">
<li>Focus on the sample using the 40x dry objective.</li>
<li>Switch to the ‘Live’ view on the CCD and adjust the exposure setting for the 40x dry
objective.</li>
<li>Rotate the dry objective out of the way and place a drop of oil on the slide and
rotate the 40x oil objective into place. Re-focus and observe the sample through
the eyepieces.</li>
<li>Do you see more detail with the one of the lenses? Explain why this is so.</li>
<li>Switch back to the ‘Live’ view on the CCD. What is the effect of NA on the CCD
exposure as well as image contrast?</li>
</ol>

</ol>

<h1>Questions & Tasks</h1>
<ol>
<li>Why do we use Kohler illumation? What is the major advantage of this illumination setup?</li>
<li>Briefly describe the difference between darkfield and brightfield and phase contrast
imaging.</li>
<li>How does the NA affect lateral (X-Y) resolution on images? Is there any effect along the Z
(focusing) axis?</li>
<li>Describe why oil immersion lenses produce sharper images as well as some of the things to
be cautious about when using immersion optics.</li>
<li>Calibrate the images you acquired and generate image stacks using the method described in
Appendix 3.</li>
<li>Calculate the cytoplasmic streaming rate in the onion cell using the method described in
Appendix 4 and compare with the value reported in the Stromgren-Allen & Brown paper.</li>
<li>You can create a movie of your cytoplasmic streaming data sets by saving your stack as an
AVI file within ImageJ (select ‘File’ -> ‘Save As’ -> ‘AVI’). Using a framerate of ~ 2 will show
the particles moving in real time. Also create movies running at 8 fps (4x real time) to see a
much more dynamic version of the streaming.</li>
</ol>

<h1>Appendix 1 - Background on Microscopy & Optics</h1>
<p>The most basic function of any microscope is to magnify an image and enhance the resolution
visible with the naked eye. In general, increasing magnification also increases the resolution
when the optical system being used has been designed and aligned properly. Before getting
into specifics, let’s review some fundamentals of how lenses and light form images.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig11_rays.png|500px|border|center]]
<b>Figure 1 -</b> Imaging properties of a single thin lens. The location and size of the image produced can be calculated
via the lens maker’s formula and the magnification ratio.
<br clear=right>
</p>
</td></table>

<p>The simplest way to produce a magnified image of an object is with a single lens, such as a
magnifying lens. Figure 1 shows a diagram that most of you should have seen at least once in
your university career, it is of a single ‘ideal’ thin lens and shows how light from the top of an
object is translated to an image plane by a lens with focal length ‘f’. Using the lens maker’s and
magnification formulas in the figure, the location and size of an object can be calculated if the
focal length and object size/position are known. When an object lies at a distance of 2f from the
lens, the image is formed at -2f and the magnification factor is -1 (ie, the image is inverted). As
the lens is moved closer to the object (decreasing d<sub>obj</sub>), the image distance and magnification
increase.</p>
<p>It is this image forming principle that is the basis for optical microscopes. The most basic form
of microscope, the compound microscope shown in Figure 2, contains two lenses. The objective
lens which collects light from the sample and the eyepiece lens, which further magnifies the
image generated by the objective and translates it to the back of the eye, or a detector such as a
CCD camera.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig12_compound.png|500px|border|center]]
<b>Figure 2 -</b> A simple compound microscope that can be used to produce a magnified image of a specimen. The
dotted line represents the apparent image size as seen by the observer or detector.
<br clear=right>
</p>
</td></table>
<p>In reality, each of these ‘lenses’ shown in Figures 1 & 2 are made up of a number of optical
components with differing shapes and glass compositions. A single lens is typically shown in the
figures to represent a multi-element lens, as diagrams can get very complex if this is not done.
The use of multiple lenses in a compound microscope allows a larger magnification factor to be
achieved, and also allows for corrections of image distortions (aberrations) which degrade image
quality when not properly corrected. A discussion of aberration correction and optical design is
beyond the scope of this lab and most undergraduate coursework, thus we will not go into this
in depth.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig13_na.png|400px|border|center]]
<b>Figure 3 -</b> Numerical Aperture (NA) is a measurement of the collection volume of a lens. In the figure, D represents
the lens diameter, f the focal length of the lens, n the index of refraction of the surrounding medium, and θ is the
maximum collection angle of the lens.
<br clear=right>
</p>
</td></table>
<p>Along with its magnification factor, the Numerical Aperture (NA) of an objective lens is very
important in reproducing sharp features in a specimen. The NA is a measure of the cone of light
collected from an objective lens; a higher NA indicates a lens that is more efficient at light
collection and produces a smaller focused spot size, which translates into better image contrast
and resolution. A diagram of the NA measure is shown in Figure 3. Light is focused to a point
which is one focal length from the lens centre. If the lens diameter is defined as D, then the
maximum collection angle of the lens for an on-axis point is then θ = tan-1(2f/D). The NA of a
lens is defined as NA = n x sin(θ), where n is the index of refraction of the medium between the
lens and the sample.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig14_focus.png|400px|border|center]]
<b>Figure 4 -</b> Focused spot size dependence on NA. As the NA is increased the collection volume increases and the
focused spot size gets smaller. This allows higher NA objectives to see finer details with better resolution and
sensitivity.
<br clear=right>
</p>
</td></table>
<p>As the NA is increased, the size of the focused spot decreases and the optical resolution
increases (Figure 4). Strictly speaking however, the distribution of the focused spot is not
actually a ‘spot’ but an Airy pattern, named after George Biddel Airy. This pattern arises due to
diffraction from the circular aperture of the lens, and the physical implication of this is that a
single point is not imaged into another single point, but into a distribution described by the Airy
pattern (Figure 5). The diameter of the Airy pattern (D<sub>Airy</sub>) can be calculated if the NA and
wavelength of light being used are known, and typically ½ of this value (Airy Radius) is defined as
the resolution limit of the system. Objects that are > 1 Airy unit apart will be clearly resolved as
two distinct points, while two points separated by < 1 Airy unit will blend together and be
indistinguishable from one another in the final image.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig15_airy.png|400px|border|center]]
<b>Figure 5 -</b> Images of two closely spaced objects as they approach the Airy resolution limit (A-C). Optical resolution
can be defined in terms of the width of the Airy pattern; two points separated by > 1 Airy radius will be resolved as
distinct objects, while objects closer than 1 Airy radius will blend together and appear as a single object (D).
<br clear=right>
</p>
</td></table>
<p>In practice, there are some fundamental limits to the magnitude of the NA possible. In the
equation for NA, sin(θ) is maximized when θ = 90°, but it is physically impossible to achieve this
collection angle in the real world. The largest NA possible for ‘dry’ objectives is around 0.95,
which corresponds to a collection angle of θ ≈ 72°. To further increase the NA beyond 0.95 a
‘wet’ or immersion objective is used. Increasing the index of refraction (n) of the region
between the lens and the sample also increases the NA, and typical immersion fluids are water
(n ≈ 1.33) and oil (n ≈ 1.55). While increasing the NA reduces the spot size, allowing smaller
features to be resolved, it also increases the light collection efficiency, thus improving the signalto-
noise ratio in images. Immersion optics are more difficult to work with, but they offer
unmatched resolution and image quality, as we will see during this lab.</p>

<h1>Appendix 2 - Phase Contrast Alignment</h1>
<p>This section gives an overview of the steps required to properly alight the phase plates in the
condenser and objective.</p>
<ol>
<li>Place a brightly stained specimen on the stage and rotate the 10x/Ph2 objective into the
optical pathway with the condenser on the BF setting. Follow the Kohler alignment
procedure from the main part of the lab.</li>
<li>Remove the sample and place a blank microscope slide on the stage.</li>
<li>Rotate the condenser turret into the Ph2 position.</li>
<li>Remove one of the microscope eyepieces and replace it with the phase telescope (if it is not
with the microscope, it will be in the wooden box that holds the DIC condenser).</li>
<li>While looking through the phase telescope, adjust its focus (slide it in and out of the tube)
until the phase plate in the objective is in sharp focus (it will look something like Figure 1a &
1c).</li>
<li>Locate the condenser annulus centering knobs and adjust the position of the annulus until it
is overlapped with the objective phase plate (Figure 1e). Note: do not attempt to adjust the
position of the condenser annulus with the main condenser centering knobs (the ones you
used for Kohler illumination). This effort will probably not achieve the intended condenser
annulus alignment and may compromise Kohler illumination.</li>
<li>The microscope should now be properly configured for phase contrast.</li>
<li>Take your prepared cheek cell sample and place it on the microscope.
<table width=500 align=center><td>
<p align=justify>[[File:Fig16_pc_align.png|400px|border|center]]
<br clear=right>
</p>
</td></table>
</ol>

<h1>Appendix 3 - Image stack generation and spatial calibration of images </h1>
<ol>
<li>Using images acquired as well as the stage micrometer images we will now generate an
image stack, which will allow you to quickly or slowly go through the data set frame by
frame. Also, the micrometer images will be used to spatially calibrate the data set.</li>
<li>Open the folder containing your data set and highlight all of the files you wish to open.</li>
<li>Drag all of the files onto the ImageJ program box (Important: to make sure the files import
in the correct numerical order make sure your mouse is on the first file when initiating the
drag and drop to ImageJ).</li>
<li>To turn the images into a stack click the ‘Image’ menu in ImageJ and select ‘Stacks’ then
‘Images to Stack’. This should combine all the separate image windows into a single
window. Moving the slider at the bottom left or right will play the sequence as a movie, or
you can go frame by frame.</li>
<li>Open the micrometer calibration image corresponding to the objective used to acquire the
stack that was just created.</li>
<li>Using the line tool in ImageJ (see below) draw a line across the marks on the slide. The lines
on the slide are separated by 10 microns, so measure from the leading edge of one black bar
to the leading edge of another bar across the field of view.
<table width=500 align=center><td>
<p align=justify>[[File:Fig17_imagej.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Open the ‘Analyze’ ImageJ menu and select ‘Set Scale’. The following window should pop
up, and already have the length of the bar you just drew (in pixels) filled in as the first entry.
Input the known calibration distance based on the number of dashes traversed by the line
and select the units to be microns. Also, check ‘Global’ to apply this calibration to all
opened images, including your stack.
<table width=200 align=center><td>
<p align=justify>[[File:Fig18_setscale.png|200px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Save and close your stack, then repeat with the rest of your images. Record the numbers for
each objective as you only have to measure the calibration once, and then can fill in the
numbers manually in the ‘Set Scale’ dialogue.</li>
<li>If you want to create a movie of your data stack select ‘File’ -> ‘Save As’ -> ‘AVI’. Selecting a
framerate of ~ 10 frames per second will be roughly 5x real time. These files can be played
in any computer media player.</li>
</ol>

<h1>Appendix 4 - Particle tracking and calculation of particle trajectories</h1>
<ol><li>Open an image stack from your onion cell data set taken in darkfield and ensure it is
calibrated properly.</li>
<li>Scan through the movie and try and identify a particle that stays in focus for at least 10
consecutive frames and can be clearly identified in each of them.</li>
<li>You can zoom in and out using the ‘+’ and ‘-‘ keys on the keyboard. You will find it easier to
track the particles if you zoom in a bit.</li>
<li>Select the ‘Segmented Lines’ tool from ImageJ by right clicking on the line tool icon as shown
below. This will allow you to left click on the particle in each frame and ‘draw’ its trajectory
on the stack. Double clicking will end drawing of the line.
<table width=400 align=center><td>
<p align=justify>[[File:Fig19_segmented.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Using the segmented line tool click on the centre of the particle you want to track. Using
the ‘<’ and ‘>’ keys on the keyboard you can scroll through the image stack frames. Track
your particle into the next frame and mark it again with a single left click, it should have
moved 10-12 microns. Repeat until the particle drifts out of view or becomes lost, ending
with a double click to finish the line draw procedure.
<table width=400 align=center><td>
<p align=justify>[[File:Fig20_stack.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>The number of boxes in the line segment indicates how many frames you tracked the
particle for. Count the number of frames you went through and call this number <charinsert>Δ</charinsert>f.</li>
<li>Now select ‘Analyze’ and then ‘Measure’ and a text box should pop up. The last entry in this
box should tell you the length of the line you just drew in microns. Call this <charinsert>Δ</charinsert>d.</li>
<li>To get the time taken, we need to confirm the time between frames from the metadata
which μManager exported while acquiring data. In the folder containing your data set there
should be a file called metadata.txt. Open this in wordpad and do a find for the highest
frame number acquired. Ie, if you set up the multi dimensional acquisition to take 100
frames, search for ‘100’. It should look like this.
<table width=400 align=center><td>
<p align=justify>[[File:Fig21_metadata.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Now, take the elapsed time (in ms) and divide it by the number of frames acquired to get
the time scale of your data set calibrated, call this value <charinsert>Δ</charinsert>t.</li>
<li>The average speed your particle is moving at is therefore:
<table width=100 align=center><td>
<p align=justify>[[File:Fig22_formula.png|100px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Repeat this calculation on multiple particles (~10) to get an average value from your data
set.</li>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-08-21T13:28:29Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://eol.org/pages/11583/overview)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie (starring Chara in the culture tank in your BPHS4090 teaching lab). The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

Main Page/BPHS 4090/Choloplast Translocation

2012-08-10T11:47:28Z

Rlew:

<h1>Required Components</h1>
<ol>
<li>[[Media:M3_Viridis.JPG|Stock culture of <i>Eremosphaera viridis</i>]]</li>
<li>[[Media:Slide_Tools.JPG|Glass slides, cover slips and micropipette]]</li>
<li>Nikon Optiphot Microscope</li>
<li>Radiometric Probe (Model S471 Portable Optometer, UDT Instruments)</li>
<li>[[Media:CCD.JPG|Cool Snap CCD Camera and MicroManager software]]</li>
<li>[[Media:M3_Band_Pass.JPG|Blue and red band-pass filters]]</li>
</ol>

<h1>Objective</h1>
<p>To observe the wavelength dependence of light on chloroplast translocation in the acidophile green
algae <i>Eremosphaera viridis</i> as well as describing a mechanism behind this response.</p>

<h1>Introduction</h1>
<p>Electromagnetic radiation is the driving force for photosynthesis. Plants and other autotrophs convert
the energy of light into chemical energy used for synthesizing carbohydrates and other organic
compounds, which heterotrophic organisms use for energy and other nutritional requirements.
Photosynthesis is a unique process performed by plants, algae, and some species of bacteria. It consists
of a series of oxidation and reduction reactions; the basic chemical formula is shown below:
<table width=380 align=center><td>
<p align=justify>[[File:Ct_eqn0.png|340px|center]]
<br clear=right>
</p>
</td></table>
Photosynthesis would not be possible without the energy derived from light. Electromagnetic radiation
enters the earth’s atmosphere from the sun where it is harvested by photosynthetic organisms.
Wavelengths between 400 and 800 nm are used by photosynthetic organisms. The sun can be
approximated as a near perfect blackbody with a surface temperature of 5800 K described by Planck’s
Black Body Radiation Law, shown in Equation 1.1
<table width=300 align=center><td>
<p align=justify>[[File:Ct_eqn1_1.png|280px|right]]
<br clear=right>
</p>
</td></table>
Where U (λ, T) has units of J m<sup>-2</sup> s<sup>-1</sup>, h is Planck’s constant, K is Boltzmann’s Constant and c is the speed
of light. The color temperature of the sun corresponds to a peak wavelength emission of about 500nm
according to Wien’s Displacement Law (Equation 1.2)
<table width=180 align=center><td>
<p align=justify>[[File:Ct_eqn1_2.png|160px|right]]
<br clear=right>
</p>
</td></table>
Where b = 2.898 x 10<sup>-3</sup> in units of m*K.</p>

<p>The first step in the conversion of the energy of photons to chemical energy is absorption by
photosynthetic pigments, principally chlorophyll. The absorption spectrum of chlorophyll a is shown in
Figure 1.</p>

<table width=800 align=center><td>
<p align=justify>[[File:Chlorophyll_a_spectrum.png|300px|left]]
<b>Absorption Spectrum of Chlorophyll a:</b> Chlorophyll a strongly absorbs
light at 465nm and 665nm. There is minimal
absorption of UV light as well as green/yellow light
which appears in the range of 500nm-600nm. These
absorbances are for chlorophyll isolated in a
solvent of a well-defined dielectric. In the cell,
additional pigments and heterogeneous dielectric
causes much broader peaks.
<br clear=right>
</p>
</td></table>

<table align=right border=0 width=400>
<tr><td>[[File:Fluorescence_scans.png|200px|left]]</td>
<td><b>Determining excitation and emission spectra</b><br>Here is an example of absorbance/fluorescence scans of whole algal cells and acetone/water extracted chlorophyll to give you an idea about the kind of absorbance that occurs
during light irradiation of your cells.<ref>http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/BIOL_4160/Determining_Excitation_and_Emission_Spectra</ref></td></tr>
</table>

<p>Chlorophyll is located inside the thylakoid vesicles in the inner membrane of the chloroplast. After
absorbing a photon, an electron in chlorophyll transitions to an excited state. The excited state electron
(exciton) is transferred to a photosynthetic reaction center to begin the flow of electrons through the
electron transport chain, eventually to produce ATP (from a transmembrane H<sup>+</sup> gradient) and reducing
equivalents (NADP + 2e<sup>–</sup> + 2H<sup>+</sup> —> NADPH + H<sup>+</sup>). Chlorophyll, now with one less electron, is chemically
unstable and receives an electron from a water molecule (H<sub>2</sub>O). With the loss of 4 electrons from two
molecules of water, molecular oxygen (O<sub>2</sub>) is produced, as well as 4H<sup>+</sup> used for ATP synthesis. Oxygen,
the waste product of photosynthesis, is vital for heterotrophs in the process of cellular respiration.</p>

<p>As light intensity increases, so do absorption events and the generation of excited state chlorophylls,
and downstream electron transport. At a high enough light intensity, these can cause the formation of a
variety of undesirable oxidative products that can damage the photosynthetic apparatus. Examples
include triplet state chlorophyll, and various reactive oxygen species (through direct reduction of O<sub>2</sub> to
O<sub>2</sub><sup>–</sup>, and subsequent formation of H<sub>2</sub>O<sub>2</sub>). The general term photo-oxidation is used to describe this
damaging process. There are protective mechanisms to avoid oxidative damage <ref>Li, Z., Wakao, S., Fischer, B.B., Niyogi, K.K. 2009. Sensing and responding to excess light. Annual Review of Plant Biology <b>60</b>: 239–260.</ref>; one of these may be
to decrease the absorptive cross-sectional area of chloroplasts by changing their location in the cell <ref>Kasahara M., Kagawa, T., Oikawa, K., Suetsugu, N., Miyao, M., Wada, M. 2002. Chloroplast avoidance movement reduces photodamage in plants. Nature 420: 829–832</ref>. The phenomenon known as systrophe is described as the accumulation of cytoplasmic organelles
around the nucleus <ref>Weidinger, M. 1980. The inhibition of systrophe by cytochalasin B. Protoplasma <b>102</b>: 167–170.</ref><ref>Weidinger, M. 1982. The inhibition of systrophe in different organisms. Protoplasma <b>110</b>: 71–
74.</ref><ref>Weidinger, M., Ruppel, H.G. 1985. Ca<sup>2+</sup> requirement for a blue-light-induced chloroplast
translocation in <i>Eremosphaera viridis</i>. Protoplasma <b>124</b>: 184–187.</ref>. Systrophe of the chloroplasts is observed in the unicellular green algae <i>Eremosphaera viridis</i> when the cell is exposed to intensities of light greater than it would experience in
its natural environment.</p>
<p>During this experiment we will be examining chloroplast translocation in the unicellular green algae
<i>Eremosphaera viridis</i>, in particular we will be looking at the wavelength dependence of light on
systrophe. A typical cell of <i>E. viridis</i> is shown in Figure 2 during the course of a light treatment protocol.</p>

<table width=800 align=center><td>
<p align=justify>[[File:Ct_fig2.png|780px|center]]
<b>Figure 2: Examples of incipient and Complete Systrophe in <i>Eremosphaera
viridis</i></b> This particular cell was illuminated with a photon flux of 200 μmol m<sup>-2</sup> s<sup>-1</sup> using a blue bandpass filter with a peak wavelength of 441nm ± 10nm.
<br clear=right>
</p>
</td></table>

<p>From Figure 2, it can be seen that the systrophe effect is quite dramatic! At 0 minutes is how the cell
would appear in its resting state, at 20 minutes into the light treatment protocol there are signs of
chloroplast translocation, this cell would be classified as incipient systrophe since it is in the process of
undergoing systrophe. Lastly at 60 minutes into the light treatment the cell would be classified as
complete systrophe. Notice at 60 minutes the cytoplasmic strands which chloroplasts migrate along
during light treatments are clearly visible!</p>
<p>An <i>Eremosphaera viridis</i> cell has about 400 chloroplasts, based on counting of medial sections of
fluorescence images. In a normal chloroplast, there is about 9•10<sup>–13</sup> g of chlorophyll and about 6.7 • 10<sup>8</sup>
chlorophyll molecules per chloroplast. The molecular weight of chlorophyll a is about 894; its extinction
coefficient is 1.2 • 10<sup>5</sup> M<sup>–1</sup> cm<sup>–1</sup> at 430nm <ref>Lawlor, D.W. 2001. Photosynthesis. Third edition. Springer-Verlag, New York. Chapters 3 and
4.</ref>.</p>

<h1>Materials and Methods </h1>

<p>The procedure is divided into four sections:
<ol style="list-style-type:upper-latin">
<li>Preparing a sample slide</li>
<li>Microscope Set-Up and Kohler Illumination</li>
<li>Light Treatments</li>
<li>Image Processing</li>
</ol>
Please follow them in order; each section has its own set of step by step instructions.</p>

<ol style="list-style-type:upper-latin">
<h2><li>Preparing a Sample Slide</h2>
<ol>
<li>Take a new glass slide and trace the outline of the cover slip using a permanent black marker as
shown in Figure 3A.</li>
<li>Spread a thin layer of Vaseline as shown in Figure 3C & D along the outline you just traced using
the syringe (Figure 3B), this will prevent your cell sample from drying out under high light
tensities</li>
<li>Aliquot 10μL of <i>E. viridis</i> in the outline you just traced using a micropipette as shown in Figure
3E</li>
<li>Gently place the cover slip on top of the Vaseline layer, being careful not to crush your cell
sample</li>
<li>Now the slide is ready to be used for light treatments (Figure 3F).</li>
<table width=600 align=center><td>
<p align=justify>[[File:Ct_fig3.png|580px|center]]
<b>Figure 3</b>
<br clear=right>
</p>
</td></table>
</ol>
</li>
<h2><li>Microscope Set-up and Kohler Illumination:</h2>
<ol>
<li>This experiment will be performed using bright-field microscopy. Ensure the ring on the
condenser diaphragm is set at 0 (not PH 1 or PH 2), because the phase ring at the PH1 or PH2
setting attenuates the light.</li>
<li>Once your specimen slide is prepared, bring it to the microscope and focus on the specimen
under the x10 objective.</li>
<li>Next close the field diaphragm all the way shut so you can see the edges, they may appear
blurry.</li>
<li>Use the condenser focus knob to bring the edges of the field diaphragm into the best possible
focus.</li>
<li>Next use the condenser-centering screws to bring the closed field diaphragm into the center of
the field of view.</li>
<li>Then open the field diaphragm such that it is slightly larger than the field of view.</li>
<li>You are now set up for Kohler illumination.</li>
</ol>
</li>
<h2><li>Light Treatments</h2>
<p>The wavelength dependence of light on chloroplast translocation will be investigated. From the
absorption spectrum of chlorophyll shown in Figure 1, it is known the chlorophyll strongly absorbs both
blue and red light, thus for the wavelength experiments we will be using blue and red filters band-pass
filters, their transmissions are shown in Figure 4. The blue band-pass filter emits light at a peak
wavelength of 441nm ± 10nm. The red filter is a band-pass filter with a tail at longer wavelengths. This
filter has a peak wavelength of 623nm ± 45nm.</p>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig4.png|450px|left]]
<b>Figure 4: Emission Spectra of Band-pass Filters:</b>The blue bandpass
filter was fit to a
Gaussian function and the
red band-pass filter was fit
to a Lorentzian Function to
account for the long tail
end. Both fit functions are
shown in black.
<br clear=right>
</p>
</td></table>

<p>The cells will be illuminated with both blue and red light to determine the wavelength dependence of
light on chloroplast translocation as described in the following outline:</p>
<ol>
<li>Take the blue band-pass filter labelled 441nm and place it above the light source under the
condenser diaphragm.</li>
<li>Double click on the Desktop folder called <b>BPHS_4090_Viridis_Systrophe</b>, and then open the
Excel spreadsheet called <b>“Photon_Flux_Calculator.xls”</b>. This spreadsheet is already
programmed to do all the difficult calculations for you to save you time in the lab!</li>
<li>With the blue filter in place turn on the radiometer, use the probe to measure the output power
from the light source; this is done by removing the sample slide and placing the probe in the
light path where the objective is. You may have to remove one of the eyepieces to get the
probe in the light path, do this carefully and ensure you leave the x10 objective (labelled Fluor
10) in place, since this will be used for the light treatments. Ensure the probe is collecting all the
light. You should measure a power reading in microwatts; enter this into the spreadsheet to get
a photon flux as an output in units of μmol m<sup>-2</sup> s<sup>-1</sup> as shown in Figure 5.</li>

<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig5.png|650px|left]]
<b>Figure 5:</b> The flux calculator has separate sections for the red and blue bandpass
filters. The cells will feel different energies corresponding to the different wavelengths of light
according to the Planck relationship <i>E = hc/λ </i>.
<br clear=right>
</p>
</td></table>

<li>For the wavelength experiment a photon flux of around <b> 200μmol m<sup>-2</sup> s<sup>-1</sup></b> is ideal to observe the
effect of chloroplast translocation. Adjust the voltage dial knob on the microscope until you get
a power reading which gives you a photon flux close to this value,<b>once you do so, record your
input power.</b></li>
<li>Next place the cell sample back in the field of view, drop the neutral density filter labelled ND 2
in the light path to avoid pre-treating the cells with light. You’ll have to adjust the microscope
again for Kohler Illumination, but it shouldn’t take much adjusting at this point.</li>
<li>Move the microscope stage to capture as large of a cell population as possible, try to aim for 20
or more cells if possible by looking through the eyepiece as you move the stage.</li>
<li>Ensure your cell sample is in good focus, you’re now ready to make a movie of chloroplast
translocation.</li>
</ol>

<h2><li>Making a Movie of Chloroplast Translocation:</h2>
<ol>
<li>Turn on the CoolSnap CCD camera.</li>
<li>Open MicroManager 1.3 by double clicking the icon on the desktop, you’ll notice the program
ImageJ (Figure 6) opens as well, this will be used later for image processing.
Figure 6 – MicroManager Software</li>
<table width=600 align=center><td>
<p align=justify>[[File:Ct_fig6.png|550px|left]]
<b>Figure 6:</b> MicroManager Software.
<br clear=right>
</p>
</td></table>
<li>Click on Live Feed and turn the eyepiece mounting block about 45 degrees to the left, this allows
light to pass through to the CCD camera and will project a live image onto the computer
monitor.</li>
<li>Next lift up the ND2 filter and adjust the exposure time, start by setting it to 1ms, you’ll also
notice the cells on the screen are out of focus, use the fine focus adjustment until you get crisp
edges along the periphery of the cell.</li>
<li>Adjust the exposure until you get a live image that isn’t oversaturated (Figure 7).</li>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig7.png|650px|left]]
<b>Figure 7:</b> MicroManager Live Feed.
<br clear=right>
</p>
</td></table>
<li>Due to the limited field of view, you’ll only be able to get about 1 cell in the live feed. Move the
microscope stage slowly (since the response from the CCD is slightly delayed) until you get 1 cell
in the field of view as shown in Figure 7.</li>
<li>Click on “Multi-D acq.” Then set the number to 360 and interval to 10 seconds, and then hit
acquire (Figure 8). This tells the program to take an image every 10 seconds; this will eventually
give you a movie 60 minutes long, which should allow you to see some significant chloroplast
translocation as shown in Figure 2.</li>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig8.png|450px|center]]
<b>Figure 8:</b> Multi-Dimensional Acquisition.
<br clear=right>
</p>
</td></table>
<li>Once finished click on the save icon. Save your data in the directory
<b>C:\BPHS4090_Viridis_Systrophe, label your data by your name, date and title of work.</b> This
will create a file folder with all the images you acquired.</li>
<table width=300 align=center><td>
<p align=justify>[[File:Ct_fig9.png|250px|center]]
<b>Figure 9:</b> Sequence Options.
<br clear=right>
</p>
</td></table>
<li>Next you’ll be using ImageJ. Click “File->Import->Image
Sequence”, double click on your file folder and select
one of the images and press Open. Another window
will open called “Sequence Options” (Figure 9), it will
verify the number of images you have and open them
as a stack, make sure the box titled “Sort Names
Numerically” is checked and click Ok.</li>
<li>This may take a little while, once all the images are
loaded, use the mouse cursor icon to make a box
around your cell and click on “Image->Crop”. Every
image you imported will be cropped.</li>
<li>Next click on “Image->Adjust->Brightness/Contrast”,
another window called “B&C” will open; adjust the
settings to ensure your images look the best they can.
Once happy with the results click on Set, a new window
will open, make sure the “Propagate to all open
images” box is checked and click OK (Figure 10). This
will adjust the brightness and contrast for every image
in your imported stack.</li>

<table width=400 align=center><td>
<p align=justify>[[File:Ct_fig10.png|350px|center]]
<b>Figure 10:</b> Adjusting Image Brightness and Contrast.
<br clear=right>
</p>
</td></table>
<li>Once your image is adjusted for proper brightness and contrast select “File->Save as->AVI”,
select a frame rate of about 10fps, this will make a movie 36 seconds long assuming you took
360 frames. A movie at this speed will dramatically show chloroplast translocation assuming the
one cell you chose to view undergoes the effect. Save the AVI in the directory
<b>C:\BPHS4090_Viridis_Systrophe by your name, date and title of work.</b></li>
<li><b>Copy your movie and the file called “Viridis_Systrophe_Spectrum.xls” onto a USB Key, this
spreadsheet will be used for one of the questions. You can find the file in the desktop folder
called “BPHS_4090_Viridis_Systrophe”.</b></li>
<li>Look at your cell sample through the field of view and record how many cells are incipient
systrophe and how many are complete systrophe using the criteria in Figure 2.</li>
<li>Once this is done discard your cell sample slide in the bin labelled “Sharps”</li>
<li>Next prepare a new sample slide and repeat the above procedure using the red filter labelled
623nm. DO NOT make a movie using the red filter; record the total number of cells in the field
of view and illuminate the cells for an hour. After the light treatment record the number of
incipient and complete systrophe. Be sure to adjust your power to get the desired photon flux
of 200 μmol m<sup>-2</sup> s<sup>-1</sup> and ensure your microscope is set up for Kohler illumination.</li>
</ol>
</ol>

<h1> Questions/Write-up </h1>

<p>Answer the following questions in your lab notebook.</p>
<ol>
<li>An additional experiment was performed [8] by illuminating only half the cell at a time. The
results of this experiment are shown below:
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig11.png|700px|center]]
<b>Figure 11:</b> Half Cell Illuminations: This particular cell was illuminated under broad band-pass
irradiation with a photon flux of <b>1500μmol m<sup>-2</sup> s<sup>-1</sup>.</b> After an hour of irradiation, chloroplasts have
vacated the illuminated area.
<br clear=right>
</p>
</td></table>
<p>What can you conclude about these results? Is chloroplast translocation a protective
mechanism for the cell or rather an avoidance mechanism? Explain in a few sentences.</p></li>

<li>Imagine you had a source of UV light available and you wished to perform an additional
experiment by illuminating the cells with UV light as you did earlier with the blue and red bandpass
filters. Assume your UV light source emits at a peak wavelength of 250nm ± 10nm. Taking
into account the absorption spectra of chlorophyll shown in Figure 1 and that DNA absorbs light
at 260nm what would you expect to happen?</li>

<li>During this lab we examined the wavelength dependence of light on systrophe, however other
experiments [8] demonstrated there is also an intensity dependence as shown in the Figure
below:</li>

<table width=500 align=center><td>
<p align=center>[[File:Ct_fig13.png|500px|center]]
<b>Figure 13:</b> Intensity Dependence of Light on Incipient and Complete Systrophe.
<br clear=right>
</p>
</td></table>

<p>At a photon flux of around <b>2000μmol m<sup>-2</sup> s<sup>-1</sup></b> there is a dramatic increase in the number of
incipient systrophe. Remember incipient means the cells are in the process of undergoing
chloroplast translocation. The photon flux we receive from the sun is around <b>2000μmol m<sup>-2</sup> s<sup>-1</sup></b>.
What does this imply?</p></li>

<li>Photosynthetic organisms use wavelengths between 400nm and 800nm for photosynthesis.
From Figure 1, it can be seen that chlorophyll strongly absorbs blue and red light. Taking this
into account and your answers to questions 1 and 2 what role do you think systrophe plays in
photosynthesis? Are they related? Please discuss in a few sentences providing insights into
your thinking process.</li>
</ol>

<p>For the formal write-up for this experiment, briefly describe the purpose of the experiment, the
experimental set-up and the physics behind the biological process we’re observing, this will be your
introduction. You should type up your results into a table comparing the illumination trials using red
and blue filters. Provide a brief discussion interpreting your results. In addition you need to provide
answers to the following questions.</p>
<ol>
<li>The effect of systrophe is thought to be a defence mechanism for the cell by decreasing the
absorptive cross sectional area of chloroplasts within the cell. We can test this hypothesis by
using a simple mathematical model to represent the cell. Assume each cell of E. viridis in its
resting state is a sphere with a shell of chloroplasts along the periphery of the cell. In reality
chloroplasts are also distributed radially along the cytoplasmic strands within the cell (analogous
to the spokes of a bicycle wheel); however for simplicity we will only consider the outer shell of
chloroplasts. After the cell undergoes systrophe, the chloroplasts surround the nucleus in a
thicker shell. In the table below some data are provided of cellular dimensions for 3 arbitrary
cells before and after systrophe using time lapsed images <ref>Web-published: http://www.yorku.ca/planters/student_reports/viridis_systrophe.pdf (accessed
9 September 2010).</ref>.

<table width=500 align=center>
<tr><td><p align=center><b>Table 1:</b> Cellular Dimensions before and after Systrophe.</p></td></tr>

<tr><td><table width=500 align=center border>
<tr><td></td><td>Cell 1</td><td>Cell 2</td><td>Cell 3</td></tr>
<tr><td>Cell Diameter (μm)</td><td>122.5</td><td>108.5</td><td>130.5</td></tr>
<tr><td>Systrophe Diameter (μm)</td><td>80.1</td><td>64.7</td><td>103.8</td></tr>
<tr><td>Nucleus Diameter (μm)</td><td>41.8</td><td>45.8</td><td>61.3</td></tr>
<tr><td>Thickness of Chloroplast Shell
(Before Systrophe) (μm)</td><td>4.6</td><td>5.5</td><td>5.3</td></tr>
</table>

</td></tr></table>

<p>Use the data in Table 1 as well as the physical properties of the cells included in the
introductions and the Beer-Lambert Law (Equation 1.3) to fill in the table below:</p>
<table width=150 align=center><td>
<p align=justify>[[File:Ct_eqn1_3.png|150px|center]]
<br clear=right>
</p>
</td></table>
<p>Where I is the transmitted intensity of light passing through the cell, Io is the initial intensity your
light source, ε is the extinction coefficient of chlorophyll, C is the concentration of chlorophyll
and l is path length which the light passes through.</p>
<table width=700 align=center><td>
<p align=center><b>Table 2: </b> Absorptive Properties of Cells before and after Systrophe.
[[File:Ct_tab2.png|650px|center]]
<br clear=right>
</p>
</td></table>

<p>Based on your results in Table 2, does systrophe provide a protective mechanism for the cell by
decreasing the absorptive cross sectional area? Why would an increase in absorption
potentially be damaging for the cell? Consider both absorption and cross sectional area.</p>
</li>

<li>Open the spreadsheet titled <b>“Viridis_Systrophe_Spectrum.xls”</b>; this is an emission spectra of
the tungsten halogen bulb used for illuminating the cell. Calculate the total energy the cells
would receive from the whole spectrum of light emitted from this source by normalization. You
can use the spreadsheet to do your calculations and may wish to attach it to your lab report.</li>
<li>Below is the emission spectrum from the tungsten halogen bulb used for illuminations. From
the Planck Blackbody Distribution Law derive Wien’s Displacement Law which relates the colour
temperature of a blackbody to the peak wavelength it emits at. If we treat tungsten as a
blackbody, what is the color temperature associated with the peak wavelength observed in
Figure 12? Does this make sense considering the melting point of tungsten is about 3350K?
What correction do we need to take into account when deriving Wien’s Displacement Law from
Planck’s Blackbody Distribution Law to account for the unusual high melting point implied from
Figure 12?</li>

<table width=500 align=center><td>
<p align=center>[[File:Ct_fig12.png|500px|center]]
<b>Figure 12:</b> Tungsten Halogen Emission Spectrum.
<br clear=right>
</p>
</td></table>

<li>During systrophe chloroplasts are translated across the cell by molecular motors. It isn’t known
exactly which molecular motors are responsible for chloroplast translocation, however it should
be noted chloroplast translocation can be inhibited when cells of <i>E. viridis</i> are treated with 2,4 –
Dinitrophenol (DNP) [4]. The structure of DNP is shown in Figure 14.

<table width=400 align=center><td>
<p align=center>[[File:Ct_fig14.png|200px|center]]
<b>Figure 14:</b> Structure of 2,4 Dinitrophenol.
<br clear=right>
</p>
</td></table>
DNP acts as an uncoupling agent by disrupting the proton gradient the cell uses to create ATP.
Any energy the cell would have gained from ATP production is lost as heat. ATP provides the
energy for molecular motors to shuttle organelles across the cell. Based on the structure of
DNP, taking into account hydrophilicity and hydrophobicity and any partial charges on the
molecule explain how DNP inhibits chloroplast translocation. Does this provide any insight into
the energy mechanisms involved during systrophe?
</li>

<h1>References</h1>
<References/>

Main Page/BPHS 4090/Choloplast Translocation

2012-08-10T11:39:05Z

Rlew:

<h1>Required Components</h1>
<ol>
<li>[[Media:M3_Viridis.JPG|Stock culture of <i>Eremosphaera viridis</i>]]</li>
<li>[[Media:Slide_Tools.JPG|Glass slides, cover slips and micropipette]]</li>
<li>Nikon Optiphot Microscope</li>
<li>Radiometric Probe (Model S471 Portable Optometer, UDT Instruments)</li>
<li>[[Media:CCD.JPG|Cool Snap CCD Camera and MicroManager software]]</li>
<li>[[Media:M3_Band_Pass.JPG|Blue and red band-pass filters]]</li>
</ol>

<h1>Objective</h1>
<p>To observe the wavelength dependence of light on chloroplast translocation in the acidophile green
algae <i>Eremosphaera viridis</i> as well as describing a mechanism behind this response.</p>

<h1>Introduction</h1>
<p>Electromagnetic radiation is the driving force for photosynthesis. Plants and other autotrophs convert
the energy of light into chemical energy used for synthesizing carbohydrates and other organic
compounds, which heterotrophic organisms use for energy and other nutritional requirements.
Photosynthesis is a unique process performed by plants, algae, and some species of bacteria. It consists
of a series of oxidation and reduction reactions; the basic chemical formula is shown below:
<table width=380 align=center><td>
<p align=justify>[[File:Ct_eqn0.png|340px|center]]
<br clear=right>
</p>
</td></table>
Photosynthesis would not be possible without the energy derived from light. Electromagnetic radiation
enters the earth’s atmosphere from the sun where it is harvested by photosynthetic organisms.
Wavelengths between 400 and 800 nm are used by photosynthetic organisms. The sun can be
approximated as a near perfect blackbody with a surface temperature of 5800 K described by Planck’s
Black Body Radiation Law, shown in Equation 1.1
<table width=300 align=center><td>
<p align=justify>[[File:Ct_eqn1_1.png|280px|right]]
<br clear=right>
</p>
</td></table>
Where U (λ, T) has units of J m<sup>-2</sup> s<sup>-1</sup>, h is Planck’s constant, K is Boltzmann’s Constant and c is the speed
of light. The color temperature of the sun corresponds to a peak wavelength emission of about 500nm
according to Wien’s Displacement Law (Equation 1.2)
<table width=180 align=center><td>
<p align=justify>[[File:Ct_eqn1_2.png|160px|right]]
<br clear=right>
</p>
</td></table>
Where b = 2.898 x 10<sup>-3</sup> in units of m*K.</p>

<p>The first step in the conversion of the energy of photons to chemical energy is absorption by
photosynthetic pigments, principally chlorophyll. The absorption spectrum of chlorophyll a is shown in
Figure 1 <ref>http://www.bio.davidson.edu/courses/Bio111/Bio111LabMan/lab1fig3.gif</ref>.</p>

<table width=800 align=center><td>
<p align=justify>[[File:Chlorophyll_a_spectrum.png|300px|left]]
<b>Absorption Spectrum of Chlorophyll a:</b> Chlorophyll a strongly absorbs
light at 465nm and 665nm. There is minimal
absorption of UV light as well as green/yellow light
which appears in the range of 500nm-600nm. These
absorbances are for chlorophyll isolated in a
solvent of a well-defined dielectric. In the cell,
additional pigments and heterogeneous dielectric
causes much broader peaks.
<br clear=right>
</p>
</td></table>

<table align=right border=0 width=400>
<tr><td>[[File:Fluorescence_scans.png|200px|left]]</td>
<td><b>Determining excitation and emission spectra</b><br>Here is an example of absorbance/fluorescence scans of whole algal cells and acetone/water extracted chlorophyll to give you an idea about the kind of absorbance that occurs
during light irradiation of your cells.</td></tr>
</table>

<p>Chlorophyll is located inside the thylakoid vesicles in the inner membrane of the chloroplast. After
absorbing a photon, an electron in chlorophyll transitions to an excited state. The excited state electron
(exciton) is transferred to a photosynthetic reaction center to begin the flow of electrons through the
electron transport chain, eventually to produce ATP (from a transmembrane H<sup>+</sup> gradient) and reducing
equivalents (NADP + 2e<sup>–</sup> + 2H<sup>+</sup> —> NADPH + H<sup>+</sup>). Chlorophyll, now with one less electron, is chemically
unstable and receives an electron from a water molecule (H<sub>2</sub>O). With the loss of 4 electrons from two
molecules of water, molecular oxygen (O<sub>2</sub>) is produced, as well as 4H<sup>+</sup> used for ATP synthesis. Oxygen,
the waste product of photosynthesis, is vital for heterotrophs in the process of cellular respiration.</p>

<p>As light intensity increases, so do absorption events and the generation of excited state chlorophylls,
and downstream electron transport. At a high enough light intensity, these can cause the formation of a
variety of undesirable oxidative products that can damage the photosynthetic apparatus. Examples
include triplet state chlorophyll, and various reactive oxygen species (through direct reduction of O<sub>2</sub> to
O<sub>2</sub><sup>–</sup>, and subsequent formation of H<sub>2</sub>O<sub>2</sub>). The general term photo-oxidation is used to describe this
damaging process. There are protective mechanisms to avoid oxidative damage <ref>Li, Z., Wakao, S., Fischer, B.B., Niyogi, K.K. 2009. Sensing and responding to excess light. Annual Review of Plant Biology <b>60</b>: 239–260.</ref>; one of these may be
to decrease the absorptive cross-sectional area of chloroplasts by changing their location in the cell <ref>Kasahara M., Kagawa, T., Oikawa, K., Suetsugu, N., Miyao, M., Wada, M. 2002. Chloroplast avoidance movement reduces photodamage in plants. Nature 420: 829–832</ref>. The phenomenon known as systrophe is described as the accumulation of cytoplasmic organelles
around the nucleus <ref>Weidinger, M. 1980. The inhibition of systrophe by cytochalasin B. Protoplasma <b>102</b>: 167–170.</ref><ref>Weidinger, M. 1982. The inhibition of systrophe in different organisms. Protoplasma <b>110</b>: 71–
74.</ref><ref>Weidinger, M., Ruppel, H.G. 1985. Ca<sup>2+</sup> requirement for a blue-light-induced chloroplast
translocation in <i>Eremosphaera viridis</i>. Protoplasma <b>124</b>: 184–187.</ref>. Systrophe of the chloroplasts is observed in the unicellular green algae <i>Eremosphaera viridis</i> when the cell is exposed to intensities of light greater than it would experience in
its natural environment.</p>
<p>During this experiment we will be examining chloroplast translocation in the unicellular green algae
<i>Eremosphaera viridis</i>, in particular we will be looking at the wavelength dependence of light on
systrophe. A typical cell of <i>E. viridis</i> is shown in Figure 2 during the course of a light treatment protocol.</p>

<table width=800 align=center><td>
<p align=justify>[[File:Ct_fig2.png|780px|center]]
<b>Figure 2: Examples of incipient and Complete Systrophe in <i>Eremosphaera
viridis</i></b> This particular cell was illuminated with a photon flux of 200 μmol m<sup>-2</sup> s<sup>-1</sup> using a blue bandpass filter with a peak wavelength of 441nm ± 10nm.
<br clear=right>
</p>
</td></table>

<p>From Figure 2, it can be seen that the systrophe effect is quite dramatic! At 0 minutes is how the cell
would appear in its resting state, at 20 minutes into the light treatment protocol there are signs of
chloroplast translocation, this cell would be classified as incipient systrophe since it is in the process of
undergoing systrophe. Lastly at 60 minutes into the light treatment the cell would be classified as
complete systrophe. Notice at 60 minutes the cytoplasmic strands which chloroplasts migrate along
during light treatments are clearly visible!</p>
<p>An <i>Eremosphaera viridis</i> cell has about 400 chloroplasts, based on counting of medial sections of
fluorescence images. In a normal chloroplast, there is about 9•10<sup>–13</sup> g of chlorophyll and about 6.7 • 10<sup>8</sup>
chlorophyll molecules per chloroplast. The molecular weight of chlorophyll a is about 894; its extinction
coefficient is 1.2 • 10<sup>5</sup> M<sup>–1</sup> cm<sup>–1</sup> at 430nm <ref>Lawlor, D.W. 2001. Photosynthesis. Third edition. Springer-Verlag, New York. Chapters 3 and
4.</ref>.</p>

<h1>Materials and Methods </h1>

<p>The procedure is divided into four sections:
<ol style="list-style-type:upper-latin">
<li>Preparing a sample slide</li>
<li>Microscope Set-Up and Kohler Illumination</li>
<li>Light Treatments</li>
<li>Image Processing</li>
</ol>
Please follow them in order; each section has its own set of step by step instructions.</p>

<ol style="list-style-type:upper-latin">
<h2><li>Preparing a Sample Slide</h2>
<ol>
<li>Take a new glass slide and trace the outline of the cover slip using a permanent black marker as
shown in Figure 3A.</li>
<li>Spread a thin layer of Vaseline as shown in Figure 3C & D along the outline you just traced using
the syringe (Figure 3B), this will prevent your cell sample from drying out under high light
tensities</li>
<li>Aliquot 10μL of <i>E. viridis</i> in the outline you just traced using a micropipette as shown in Figure
3E</li>
<li>Gently place the cover slip on top of the Vaseline layer, being careful not to crush your cell
sample</li>
<li>Now the slide is ready to be used for light treatments (Figure 3F).</li>
<table width=600 align=center><td>
<p align=justify>[[File:Ct_fig3.png|580px|center]]
<b>Figure 3</b>
<br clear=right>
</p>
</td></table>
</ol>
</li>
<h2><li>Microscope Set-up and Kohler Illumination:</h2>
<ol>
<li>This experiment will be performed using bright-field microscopy. Ensure the ring on the
condenser diaphragm is set at 0 (not PH 1 or PH 2), because the phase ring at the PH1 or PH2
setting attenuates the light.</li>
<li>Once your specimen slide is prepared, bring it to the microscope and focus on the specimen
under the x10 objective.</li>
<li>Next close the field diaphragm all the way shut so you can see the edges, they may appear
blurry.</li>
<li>Use the condenser focus knob to bring the edges of the field diaphragm into the best possible
focus.</li>
<li>Next use the condenser-centering screws to bring the closed field diaphragm into the center of
the field of view.</li>
<li>Then open the field diaphragm such that it is slightly larger than the field of view.</li>
<li>You are now set up for Kohler illumination.</li>
</ol>
</li>
<h2><li>Light Treatments</h2>
<p>The wavelength dependence of light on chloroplast translocation will be investigated. From the
absorption spectrum of chlorophyll shown in Figure 1, it is known the chlorophyll strongly absorbs both
blue and red light, thus for the wavelength experiments we will be using blue and red filters band-pass
filters, their transmissions are shown in Figure 4. The blue band-pass filter emits light at a peak
wavelength of 441nm ± 10nm. The red filter is a band-pass filter with a tail at longer wavelengths. This
filter has a peak wavelength of 623nm ± 45nm.</p>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig4.png|450px|left]]
<b>Figure 4: Emission Spectra of Band-pass Filters:</b>The blue bandpass
filter was fit to a
Gaussian function and the
red band-pass filter was fit
to a Lorentzian Function to
account for the long tail
end. Both fit functions are
shown in black.
<br clear=right>
</p>
</td></table>

<p>The cells will be illuminated with both blue and red light to determine the wavelength dependence of
light on chloroplast translocation as described in the following outline:</p>
<ol>
<li>Take the blue band-pass filter labelled 441nm and place it above the light source under the
condenser diaphragm.</li>
<li>Double click on the Desktop folder called <b>BPHS_4090_Viridis_Systrophe</b>, and then open the
Excel spreadsheet called <b>“Photon_Flux_Calculator.xls”</b>. This spreadsheet is already
programmed to do all the difficult calculations for you to save you time in the lab!</li>
<li>With the blue filter in place turn on the radiometer, use the probe to measure the output power
from the light source; this is done by removing the sample slide and placing the probe in the
light path where the objective is. You may have to remove one of the eyepieces to get the
probe in the light path, do this carefully and ensure you leave the x10 objective (labelled Fluor
10) in place, since this will be used for the light treatments. Ensure the probe is collecting all the
light. You should measure a power reading in microwatts; enter this into the spreadsheet to get
a photon flux as an output in units of μmol m<sup>-2</sup> s<sup>-1</sup> as shown in Figure 5.</li>

<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig5.png|650px|left]]
<b>Figure 5:</b> The flux calculator has separate sections for the red and blue bandpass
filters. The cells will feel different energies corresponding to the different wavelengths of light
according to the Planck relationship <i>E = hc/λ </i>.
<br clear=right>
</p>
</td></table>

<li>For the wavelength experiment a photon flux of around <b> 200μmol m<sup>-2</sup> s<sup>-1</sup></b> is ideal to observe the
effect of chloroplast translocation. Adjust the voltage dial knob on the microscope until you get
a power reading which gives you a photon flux close to this value,<b>once you do so, record your
input power.</b></li>
<li>Next place the cell sample back in the field of view, drop the neutral density filter labelled ND 2
in the light path to avoid pre-treating the cells with light. You’ll have to adjust the microscope
again for Kohler Illumination, but it shouldn’t take much adjusting at this point.</li>
<li>Move the microscope stage to capture as large of a cell population as possible, try to aim for 20
or more cells if possible by looking through the eyepiece as you move the stage.</li>
<li>Ensure your cell sample is in good focus, you’re now ready to make a movie of chloroplast
translocation.</li>
</ol>

<h2><li>Making a Movie of Chloroplast Translocation:</h2>
<ol>
<li>Turn on the CoolSnap CCD camera.</li>
<li>Open MicroManager 1.3 by double clicking the icon on the desktop, you’ll notice the program
ImageJ (Figure 6) opens as well, this will be used later for image processing.
Figure 6 – MicroManager Software</li>
<table width=600 align=center><td>
<p align=justify>[[File:Ct_fig6.png|550px|left]]
<b>Figure 6:</b> MicroManager Software.
<br clear=right>
</p>
</td></table>
<li>Click on Live Feed and turn the eyepiece mounting block about 45 degrees to the left, this allows
light to pass through to the CCD camera and will project a live image onto the computer
monitor.</li>
<li>Next lift up the ND2 filter and adjust the exposure time, start by setting it to 1ms, you’ll also
notice the cells on the screen are out of focus, use the fine focus adjustment until you get crisp
edges along the periphery of the cell.</li>
<li>Adjust the exposure until you get a live image that isn’t oversaturated (Figure 7).</li>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig7.png|650px|left]]
<b>Figure 7:</b> MicroManager Live Feed.
<br clear=right>
</p>
</td></table>
<li>Due to the limited field of view, you’ll only be able to get about 1 cell in the live feed. Move the
microscope stage slowly (since the response from the CCD is slightly delayed) until you get 1 cell
in the field of view as shown in Figure 7.</li>
<li>Click on “Multi-D acq.” Then set the number to 360 and interval to 10 seconds, and then hit
acquire (Figure 8). This tells the program to take an image every 10 seconds; this will eventually
give you a movie 60 minutes long, which should allow you to see some significant chloroplast
translocation as shown in Figure 2.</li>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig8.png|450px|center]]
<b>Figure 8:</b> Multi-Dimensional Acquisition.
<br clear=right>
</p>
</td></table>
<li>Once finished click on the save icon. Save your data in the directory
<b>C:\BPHS4090_Viridis_Systrophe, label your data by your name, date and title of work.</b> This
will create a file folder with all the images you acquired.</li>
<table width=300 align=center><td>
<p align=justify>[[File:Ct_fig9.png|250px|center]]
<b>Figure 9:</b> Sequence Options.
<br clear=right>
</p>
</td></table>
<li>Next you’ll be using ImageJ. Click “File->Import->Image
Sequence”, double click on your file folder and select
one of the images and press Open. Another window
will open called “Sequence Options” (Figure 9), it will
verify the number of images you have and open them
as a stack, make sure the box titled “Sort Names
Numerically” is checked and click Ok.</li>
<li>This may take a little while, once all the images are
loaded, use the mouse cursor icon to make a box
around your cell and click on “Image->Crop”. Every
image you imported will be cropped.</li>
<li>Next click on “Image->Adjust->Brightness/Contrast”,
another window called “B&C” will open; adjust the
settings to ensure your images look the best they can.
Once happy with the results click on Set, a new window
will open, make sure the “Propagate to all open
images” box is checked and click OK (Figure 10). This
will adjust the brightness and contrast for every image
in your imported stack.</li>

<table width=400 align=center><td>
<p align=justify>[[File:Ct_fig10.png|350px|center]]
<b>Figure 10:</b> Adjusting Image Brightness and Contrast.
<br clear=right>
</p>
</td></table>
<li>Once your image is adjusted for proper brightness and contrast select “File->Save as->AVI”,
select a frame rate of about 10fps, this will make a movie 36 seconds long assuming you took
360 frames. A movie at this speed will dramatically show chloroplast translocation assuming the
one cell you chose to view undergoes the effect. Save the AVI in the directory
<b>C:\BPHS4090_Viridis_Systrophe by your name, date and title of work.</b></li>
<li><b>Copy your movie and the file called “Viridis_Systrophe_Spectrum.xls” onto a USB Key, this
spreadsheet will be used for one of the questions. You can find the file in the desktop folder
called “BPHS_4090_Viridis_Systrophe”.</b></li>
<li>Look at your cell sample through the field of view and record how many cells are incipient
systrophe and how many are complete systrophe using the criteria in Figure 2.</li>
<li>Once this is done discard your cell sample slide in the bin labelled “Sharps”</li>
<li>Next prepare a new sample slide and repeat the above procedure using the red filter labelled
623nm. DO NOT make a movie using the red filter; record the total number of cells in the field
of view and illuminate the cells for an hour. After the light treatment record the number of
incipient and complete systrophe. Be sure to adjust your power to get the desired photon flux
of 200 μmol m<sup>-2</sup> s<sup>-1</sup> and ensure your microscope is set up for Kohler illumination.</li>
</ol>
</ol>

<h1> Questions/Write-up </h1>

<p>Answer the following questions in your lab notebook.</p>
<ol>
<li>An additional experiment was performed [8] by illuminating only half the cell at a time. The
results of this experiment are shown below:
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig11.png|700px|center]]
<b>Figure 11:</b> Half Cell Illuminations: This particular cell was illuminated under broad band-pass
irradiation with a photon flux of <b>1500μmol m<sup>-2</sup> s<sup>-1</sup>.</b> After an hour of irradiation, chloroplasts have
vacated the illuminated area.
<br clear=right>
</p>
</td></table>
<p>What can you conclude about these results? Is chloroplast translocation a protective
mechanism for the cell or rather an avoidance mechanism? Explain in a few sentences.</p></li>

<li>Imagine you had a source of UV light available and you wished to perform an additional
experiment by illuminating the cells with UV light as you did earlier with the blue and red bandpass
filters. Assume your UV light source emits at a peak wavelength of 250nm ± 10nm. Taking
into account the absorption spectra of chlorophyll shown in Figure 1 and that DNA absorbs light
at 260nm what would you expect to happen?</li>

<li>During this lab we examined the wavelength dependence of light on systrophe, however other
experiments [8] demonstrated there is also an intensity dependence as shown in the Figure
below:</li>

<table width=500 align=center><td>
<p align=center>[[File:Ct_fig13.png|500px|center]]
<b>Figure 13:</b> Intensity Dependence of Light on Incipient and Complete Systrophe.
<br clear=right>
</p>
</td></table>

<p>At a photon flux of around <b>2000μmol m<sup>-2</sup> s<sup>-1</sup></b> there is a dramatic increase in the number of
incipient systrophe. Remember incipient means the cells are in the process of undergoing
chloroplast translocation. The photon flux we receive from the sun is around <b>2000μmol m<sup>-2</sup> s<sup>-1</sup></b>.
What does this imply?</p></li>

<li>Photosynthetic organisms use wavelengths between 400nm and 800nm for photosynthesis.
From Figure 1, it can be seen that chlorophyll strongly absorbs blue and red light. Taking this
into account and your answers to questions 1 and 2 what role do you think systrophe plays in
photosynthesis? Are they related? Please discuss in a few sentences providing insights into
your thinking process.</li>
</ol>

<p>For the formal write-up for this experiment, briefly describe the purpose of the experiment, the
experimental set-up and the physics behind the biological process we’re observing, this will be your
introduction. You should type up your results into a table comparing the illumination trials using red
and blue filters. Provide a brief discussion interpreting your results. In addition you need to provide
answers to the following questions.</p>
<ol>
<li>The effect of systrophe is thought to be a defence mechanism for the cell by decreasing the
absorptive cross sectional area of chloroplasts within the cell. We can test this hypothesis by
using a simple mathematical model to represent the cell. Assume each cell of E. viridis in its
resting state is a sphere with a shell of chloroplasts along the periphery of the cell. In reality
chloroplasts are also distributed radially along the cytoplasmic strands within the cell (analogous
to the spokes of a bicycle wheel); however for simplicity we will only consider the outer shell of
chloroplasts. After the cell undergoes systrophe, the chloroplasts surround the nucleus in a
thicker shell. In the table below some data are provided of cellular dimensions for 3 arbitrary
cells before and after systrophe using time lapsed images <ref>Web-published: http://www.yorku.ca/planters/student_reports/viridis_systrophe.pdf (accessed
9 September 2010).</ref>.

<table width=500 align=center>
<tr><td><p align=center><b>Table 1:</b> Cellular Dimensions before and after Systrophe.</p></td></tr>

<tr><td><table width=500 align=center border>
<tr><td></td><td>Cell 1</td><td>Cell 2</td><td>Cell 3</td></tr>
<tr><td>Cell Diameter (μm)</td><td>122.5</td><td>108.5</td><td>130.5</td></tr>
<tr><td>Systrophe Diameter (μm)</td><td>80.1</td><td>64.7</td><td>103.8</td></tr>
<tr><td>Nucleus Diameter (μm)</td><td>41.8</td><td>45.8</td><td>61.3</td></tr>
<tr><td>Thickness of Chloroplast Shell
(Before Systrophe) (μm)</td><td>4.6</td><td>5.5</td><td>5.3</td></tr>
</table>

</td></tr></table>

<p>Use the data in Table 1 as well as the physical properties of the cells included in the
introductions and the Beer-Lambert Law (Equation 1.3) to fill in the table below:</p>
<table width=150 align=center><td>
<p align=justify>[[File:Ct_eqn1_3.png|150px|center]]
<br clear=right>
</p>
</td></table>
<p>Where I is the transmitted intensity of light passing through the cell, Io is the initial intensity your
light source, ε is the extinction coefficient of chlorophyll, C is the concentration of chlorophyll
and l is path length which the light passes through.</p>
<table width=700 align=center><td>
<p align=center><b>Table 2: </b> Absorptive Properties of Cells before and after Systrophe.
[[File:Ct_tab2.png|650px|center]]
<br clear=right>
</p>
</td></table>

<p>Based on your results in Table 2, does systrophe provide a protective mechanism for the cell by
decreasing the absorptive cross sectional area? Why would an increase in absorption
potentially be damaging for the cell? Consider both absorption and cross sectional area.</p>
</li>

<li>Open the spreadsheet titled <b>“Viridis_Systrophe_Spectrum.xls”</b>; this is an emission spectra of
the tungsten halogen bulb used for illuminating the cell. Calculate the total energy the cells
would receive from the whole spectrum of light emitted from this source by normalization. You
can use the spreadsheet to do your calculations and may wish to attach it to your lab report.</li>
<li>Below is the emission spectrum from the tungsten halogen bulb used for illuminations. From
the Planck Blackbody Distribution Law derive Wien’s Displacement Law which relates the colour
temperature of a blackbody to the peak wavelength it emits at. If we treat tungsten as a
blackbody, what is the color temperature associated with the peak wavelength observed in
Figure 12? Does this make sense considering the melting point of tungsten is about 3350K?
What correction do we need to take into account when deriving Wien’s Displacement Law from
Planck’s Blackbody Distribution Law to account for the unusual high melting point implied from
Figure 12?</li>

<table width=500 align=center><td>
<p align=center>[[File:Ct_fig12.png|500px|center]]
<b>Figure 12:</b> Tungsten Halogen Emission Spectrum.
<br clear=right>
</p>
</td></table>

<li>During systrophe chloroplasts are translated across the cell by molecular motors. It isn’t known
exactly which molecular motors are responsible for chloroplast translocation, however it should
be noted chloroplast translocation can be inhibited when cells of <i>E. viridis</i> are treated with 2,4 –
Dinitrophenol (DNP) [4]. The structure of DNP is shown in Figure 14.

<table width=400 align=center><td>
<p align=center>[[File:Ct_fig14.png|200px|center]]
<b>Figure 14:</b> Structure of 2,4 Dinitrophenol.
<br clear=right>
</p>
</td></table>
DNP acts as an uncoupling agent by disrupting the proton gradient the cell uses to create ATP.
Any energy the cell would have gained from ATP production is lost as heat. ATP provides the
energy for molecular motors to shuttle organelles across the cell. Based on the structure of
DNP, taking into account hydrophilicity and hydrophobicity and any partial charges on the
molecule explain how DNP inhibits chloroplast translocation. Does this provide any insight into
the energy mechanisms involved during systrophe?
</li>

<h1>References</h1>
<References/>

Main Page/BPHS 4090/Choloplast Translocation

2012-08-10T11:36:12Z

Rlew:

<h1>Required Components</h1>
<ol>
<li>[[Media:M3_Viridis.JPG|Stock culture of <i>Eremosphaera viridis</i>]]</li>
<li>[[Media:Slide_Tools.JPG|Glass slides, cover slips and micropipette]]</li>
<li>Nikon Optiphot Microscope</li>
<li>Radiometric Probe (Model S471 Portable Optometer, UDT Instruments)</li>
<li>[[Media:CCD.JPG|Cool Snap CCD Camera and MicroManager software]]</li>
<li>[[Media:M3_Band_Pass.JPG|Blue and red band-pass filters]]</li>
</ol>

<h1>Objective</h1>
<p>To observe the wavelength dependence of light on chloroplast translocation in the acidophile green
algae <i>Eremosphaera viridis</i> as well as describing a mechanism behind this response.</p>

<h1>Introduction</h1>
<p>Electromagnetic radiation is the driving force for photosynthesis. Plants and other autotrophs convert
the energy of light into chemical energy used for synthesizing carbohydrates and other organic
compounds, which heterotrophic organisms use for energy and other nutritional requirements.
Photosynthesis is a unique process performed by plants, algae, and some species of bacteria. It consists
of a series of oxidation and reduction reactions; the basic chemical formula is shown below:
<table width=380 align=center><td>
<p align=justify>[[File:Ct_eqn0.png|340px|center]]
<br clear=right>
</p>
</td></table>
Photosynthesis would not be possible without the energy derived from light. Electromagnetic radiation
enters the earth’s atmosphere from the sun where it is harvested by photosynthetic organisms.
Wavelengths between 400 and 800 nm are used by photosynthetic organisms. The sun can be
approximated as a near perfect blackbody with a surface temperature of 5800 K described by Planck’s
Black Body Radiation Law, shown in Equation 1.1
<table width=300 align=center><td>
<p align=justify>[[File:Ct_eqn1_1.png|280px|right]]
<br clear=right>
</p>
</td></table>
Where U (λ, T) has units of J m<sup>-2</sup> s<sup>-1</sup>, h is Planck’s constant, K is Boltzmann’s Constant and c is the speed
of light. The color temperature of the sun corresponds to a peak wavelength emission of about 500nm
according to Wien’s Displacement Law (Equation 1.2)
<table width=180 align=center><td>
<p align=justify>[[File:Ct_eqn1_2.png|160px|right]]
<br clear=right>
</p>
</td></table>
Where b = 2.898 x 10<sup>-3</sup> in units of m*K.</p>

<p>The first step in the conversion of the energy of photons to chemical energy is absorption by
photosynthetic pigments, principally chlorophyll. The absorption spectrum of chlorophyll a is shown in
Figure 1 <ref>http://www.bio.davidson.edu/courses/Bio111/Bio111LabMan/lab1fig3.gif</ref>.</p>

<table width=800 align=center><td>
<p align=justify>[[File:Chlorophyll_a_spectrum.png|300px|left]]
<b>Absorption Spectrum of Chlorophyll a:</b> Chlorophyll a strongly absorbs
light at 465nm and 665nm. There is minimal
absorption of UV light as well as green/yellow light
which appears in the range of 500nm-600nm. These
absorbances are for chlorophyll isolated in a
solvent of a well-defined dielectric. In the cell,
additional pigments and heterogeneous dielectric
causes much broader peaks.
<br clear=right>
</p>
</td></table>

<table align=right border=0 width=400>
<tr><td>[[File:Fluorescence_scans.png|200px|left]]</td>
<td><b>DETERMINING EXCITATION AND EMISSION SPECTRA</b><br>Here is an example of absorbance/fluorescence scans of whole algal cells and acetone/water extracted chlorophyll to give you an idea about the kind of absorbance that occurs
during light irradiation of your cells.</td></tr>
</table>

<p>Chlorophyll is located inside the thylakoid vesicles in the inner membrane of the chloroplast. After
absorbing a photon, an electron in chlorophyll transitions to an excited state. The excited state electron
(exciton) is transferred to a photosynthetic reaction center to begin the flow of electrons through the
electron transport chain, eventually to produce ATP (from a transmembrane H<sup>+</sup> gradient) and reducing
equivalents (NADP + 2e<sup>–</sup> + 2H<sup>+</sup> —> NADPH + H<sup>+</sup>). Chlorophyll, now with one less electron, is chemically
unstable and receives an electron from a water molecule (H<sub>2</sub>O). With the loss of 4 electrons from two
molecules of water, molecular oxygen (O<sub>2</sub>) is produced, as well as 4H<sup>+</sup> used for ATP synthesis. Oxygen,
the waste product of photosynthesis, is vital for heterotrophs in the process of cellular respiration.</p>

<p>As light intensity increases, so do absorption events and the generation of excited state chlorophylls,
and downstream electron transport. At a high enough light intensity, these can cause the formation of a
variety of undesirable oxidative products that can damage the photosynthetic apparatus. Examples
include triplet state chlorophyll, and various reactive oxygen species (through direct reduction of O<sub>2</sub> to
O<sub>2</sub><sup>–</sup>, and subsequent formation of H<sub>2</sub>O<sub>2</sub>). The general term photo-oxidation is used to describe this
damaging process. There are protective mechanisms to avoid oxidative damage <ref>Li, Z., Wakao, S., Fischer, B.B., Niyogi, K.K. 2009. Sensing and responding to excess light. Annual Review of Plant Biology <b>60</b>: 239–260.</ref>; one of these may be
to decrease the absorptive cross-sectional area of chloroplasts by changing their location in the cell <ref>Kasahara M., Kagawa, T., Oikawa, K., Suetsugu, N., Miyao, M., Wada, M. 2002. Chloroplast avoidance movement reduces photodamage in plants. Nature 420: 829–832</ref>. The phenomenon known as systrophe is described as the accumulation of cytoplasmic organelles
around the nucleus <ref>Weidinger, M. 1980. The inhibition of systrophe by cytochalasin B. Protoplasma <b>102</b>: 167–170.</ref><ref>Weidinger, M. 1982. The inhibition of systrophe in different organisms. Protoplasma <b>110</b>: 71–
74.</ref><ref>Weidinger, M., Ruppel, H.G. 1985. Ca<sup>2+</sup> requirement for a blue-light-induced chloroplast
translocation in <i>Eremosphaera viridis</i>. Protoplasma <b>124</b>: 184–187.</ref>. Systrophe of the chloroplasts is observed in the unicellular green algae <i>Eremosphaera viridis</i> when the cell is exposed to intensities of light greater than it would experience in
its natural environment.</p>
<p>During this experiment we will be examining chloroplast translocation in the unicellular green algae
<i>Eremosphaera viridis</i>, in particular we will be looking at the wavelength dependence of light on
systrophe. A typical cell of <i>E. viridis</i> is shown in Figure 2 during the course of a light treatment protocol.</p>

<table width=800 align=center><td>
<p align=justify>[[File:Ct_fig2.png|780px|center]]
<b>Figure 2: Examples of incipient and Complete Systrophe in <i>Eremosphaera
viridis</i></b> This particular cell was illuminated with a photon flux of 200 μmol m<sup>-2</sup> s<sup>-1</sup> using a blue bandpass filter with a peak wavelength of 441nm ± 10nm.
<br clear=right>
</p>
</td></table>

<p>From Figure 2, it can be seen that the systrophe effect is quite dramatic! At 0 minutes is how the cell
would appear in its resting state, at 20 minutes into the light treatment protocol there are signs of
chloroplast translocation, this cell would be classified as incipient systrophe since it is in the process of
undergoing systrophe. Lastly at 60 minutes into the light treatment the cell would be classified as
complete systrophe. Notice at 60 minutes the cytoplasmic strands which chloroplasts migrate along
during light treatments are clearly visible!</p>
<p>An <i>Eremosphaera viridis</i> cell has about 400 chloroplasts, based on counting of medial sections of
fluorescence images. In a normal chloroplast, there is about 9•10<sup>–13</sup> g of chlorophyll and about 6.7 • 10<sup>8</sup>
chlorophyll molecules per chloroplast. The molecular weight of chlorophyll a is about 894; its extinction
coefficient is 1.2 • 10<sup>5</sup> M<sup>–1</sup> cm<sup>–1</sup> at 430nm <ref>Lawlor, D.W. 2001. Photosynthesis. Third edition. Springer-Verlag, New York. Chapters 3 and
4.</ref>.</p>

<h1>Materials and Methods </h1>

<p>The procedure is divided into four sections:
<ol style="list-style-type:upper-latin">
<li>Preparing a sample slide</li>
<li>Microscope Set-Up and Kohler Illumination</li>
<li>Light Treatments</li>
<li>Image Processing</li>
</ol>
Please follow them in order; each section has its own set of step by step instructions.</p>

<ol style="list-style-type:upper-latin">
<h2><li>Preparing a Sample Slide</h2>
<ol>
<li>Take a new glass slide and trace the outline of the cover slip using a permanent black marker as
shown in Figure 3A.</li>
<li>Spread a thin layer of Vaseline as shown in Figure 3C & D along the outline you just traced using
the syringe (Figure 3B), this will prevent your cell sample from drying out under high light
tensities</li>
<li>Aliquot 10μL of <i>E. viridis</i> in the outline you just traced using a micropipette as shown in Figure
3E</li>
<li>Gently place the cover slip on top of the Vaseline layer, being careful not to crush your cell
sample</li>
<li>Now the slide is ready to be used for light treatments (Figure 3F).</li>
<table width=600 align=center><td>
<p align=justify>[[File:Ct_fig3.png|580px|center]]
<b>Figure 3</b>
<br clear=right>
</p>
</td></table>
</ol>
</li>
<h2><li>Microscope Set-up and Kohler Illumination:</h2>
<ol>
<li>This experiment will be performed using bright-field microscopy. Ensure the ring on the
condenser diaphragm is set at 0 (not PH 1 or PH 2), because the phase ring at the PH1 or PH2
setting attenuates the light.</li>
<li>Once your specimen slide is prepared, bring it to the microscope and focus on the specimen
under the x10 objective.</li>
<li>Next close the field diaphragm all the way shut so you can see the edges, they may appear
blurry.</li>
<li>Use the condenser focus knob to bring the edges of the field diaphragm into the best possible
focus.</li>
<li>Next use the condenser-centering screws to bring the closed field diaphragm into the center of
the field of view.</li>
<li>Then open the field diaphragm such that it is slightly larger than the field of view.</li>
<li>You are now set up for Kohler illumination.</li>
</ol>
</li>
<h2><li>Light Treatments</h2>
<p>The wavelength dependence of light on chloroplast translocation will be investigated. From the
absorption spectrum of chlorophyll shown in Figure 1, it is known the chlorophyll strongly absorbs both
blue and red light, thus for the wavelength experiments we will be using blue and red filters band-pass
filters, their transmissions are shown in Figure 4. The blue band-pass filter emits light at a peak
wavelength of 441nm ± 10nm. The red filter is a band-pass filter with a tail at longer wavelengths. This
filter has a peak wavelength of 623nm ± 45nm.</p>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig4.png|450px|left]]
<b>Figure 4: Emission Spectra of Band-pass Filters:</b>The blue bandpass
filter was fit to a
Gaussian function and the
red band-pass filter was fit
to a Lorentzian Function to
account for the long tail
end. Both fit functions are
shown in black.
<br clear=right>
</p>
</td></table>

<p>The cells will be illuminated with both blue and red light to determine the wavelength dependence of
light on chloroplast translocation as described in the following outline:</p>
<ol>
<li>Take the blue band-pass filter labelled 441nm and place it above the light source under the
condenser diaphragm.</li>
<li>Double click on the Desktop folder called <b>BPHS_4090_Viridis_Systrophe</b>, and then open the
Excel spreadsheet called <b>“Photon_Flux_Calculator.xls”</b>. This spreadsheet is already
programmed to do all the difficult calculations for you to save you time in the lab!</li>
<li>With the blue filter in place turn on the radiometer, use the probe to measure the output power
from the light source; this is done by removing the sample slide and placing the probe in the
light path where the objective is. You may have to remove one of the eyepieces to get the
probe in the light path, do this carefully and ensure you leave the x10 objective (labelled Fluor
10) in place, since this will be used for the light treatments. Ensure the probe is collecting all the
light. You should measure a power reading in microwatts; enter this into the spreadsheet to get
a photon flux as an output in units of μmol m<sup>-2</sup> s<sup>-1</sup> as shown in Figure 5.</li>

<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig5.png|650px|left]]
<b>Figure 5:</b> The flux calculator has separate sections for the red and blue bandpass
filters. The cells will feel different energies corresponding to the different wavelengths of light
according to the Planck relationship <i>E = hc/λ </i>.
<br clear=right>
</p>
</td></table>

<li>For the wavelength experiment a photon flux of around <b> 200μmol m<sup>-2</sup> s<sup>-1</sup></b> is ideal to observe the
effect of chloroplast translocation. Adjust the voltage dial knob on the microscope until you get
a power reading which gives you a photon flux close to this value,<b>once you do so, record your
input power.</b></li>
<li>Next place the cell sample back in the field of view, drop the neutral density filter labelled ND 2
in the light path to avoid pre-treating the cells with light. You’ll have to adjust the microscope
again for Kohler Illumination, but it shouldn’t take much adjusting at this point.</li>
<li>Move the microscope stage to capture as large of a cell population as possible, try to aim for 20
or more cells if possible by looking through the eyepiece as you move the stage.</li>
<li>Ensure your cell sample is in good focus, you’re now ready to make a movie of chloroplast
translocation.</li>
</ol>

<h2><li>Making a Movie of Chloroplast Translocation:</h2>
<ol>
<li>Turn on the CoolSnap CCD camera.</li>
<li>Open MicroManager 1.3 by double clicking the icon on the desktop, you’ll notice the program
ImageJ (Figure 6) opens as well, this will be used later for image processing.
Figure 6 – MicroManager Software</li>
<table width=600 align=center><td>
<p align=justify>[[File:Ct_fig6.png|550px|left]]
<b>Figure 6:</b> MicroManager Software.
<br clear=right>
</p>
</td></table>
<li>Click on Live Feed and turn the eyepiece mounting block about 45 degrees to the left, this allows
light to pass through to the CCD camera and will project a live image onto the computer
monitor.</li>
<li>Next lift up the ND2 filter and adjust the exposure time, start by setting it to 1ms, you’ll also
notice the cells on the screen are out of focus, use the fine focus adjustment until you get crisp
edges along the periphery of the cell.</li>
<li>Adjust the exposure until you get a live image that isn’t oversaturated (Figure 7).</li>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig7.png|650px|left]]
<b>Figure 7:</b> MicroManager Live Feed.
<br clear=right>
</p>
</td></table>
<li>Due to the limited field of view, you’ll only be able to get about 1 cell in the live feed. Move the
microscope stage slowly (since the response from the CCD is slightly delayed) until you get 1 cell
in the field of view as shown in Figure 7.</li>
<li>Click on “Multi-D acq.” Then set the number to 360 and interval to 10 seconds, and then hit
acquire (Figure 8). This tells the program to take an image every 10 seconds; this will eventually
give you a movie 60 minutes long, which should allow you to see some significant chloroplast
translocation as shown in Figure 2.</li>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig8.png|450px|center]]
<b>Figure 8:</b> Multi-Dimensional Acquisition.
<br clear=right>
</p>
</td></table>
<li>Once finished click on the save icon. Save your data in the directory
<b>C:\BPHS4090_Viridis_Systrophe, label your data by your name, date and title of work.</b> This
will create a file folder with all the images you acquired.</li>
<table width=300 align=center><td>
<p align=justify>[[File:Ct_fig9.png|250px|center]]
<b>Figure 9:</b> Sequence Options.
<br clear=right>
</p>
</td></table>
<li>Next you’ll be using ImageJ. Click “File->Import->Image
Sequence”, double click on your file folder and select
one of the images and press Open. Another window
will open called “Sequence Options” (Figure 9), it will
verify the number of images you have and open them
as a stack, make sure the box titled “Sort Names
Numerically” is checked and click Ok.</li>
<li>This may take a little while, once all the images are
loaded, use the mouse cursor icon to make a box
around your cell and click on “Image->Crop”. Every
image you imported will be cropped.</li>
<li>Next click on “Image->Adjust->Brightness/Contrast”,
another window called “B&C” will open; adjust the
settings to ensure your images look the best they can.
Once happy with the results click on Set, a new window
will open, make sure the “Propagate to all open
images” box is checked and click OK (Figure 10). This
will adjust the brightness and contrast for every image
in your imported stack.</li>

<table width=400 align=center><td>
<p align=justify>[[File:Ct_fig10.png|350px|center]]
<b>Figure 10:</b> Adjusting Image Brightness and Contrast.
<br clear=right>
</p>
</td></table>
<li>Once your image is adjusted for proper brightness and contrast select “File->Save as->AVI”,
select a frame rate of about 10fps, this will make a movie 36 seconds long assuming you took
360 frames. A movie at this speed will dramatically show chloroplast translocation assuming the
one cell you chose to view undergoes the effect. Save the AVI in the directory
<b>C:\BPHS4090_Viridis_Systrophe by your name, date and title of work.</b></li>
<li><b>Copy your movie and the file called “Viridis_Systrophe_Spectrum.xls” onto a USB Key, this
spreadsheet will be used for one of the questions. You can find the file in the desktop folder
called “BPHS_4090_Viridis_Systrophe”.</b></li>
<li>Look at your cell sample through the field of view and record how many cells are incipient
systrophe and how many are complete systrophe using the criteria in Figure 2.</li>
<li>Once this is done discard your cell sample slide in the bin labelled “Sharps”</li>
<li>Next prepare a new sample slide and repeat the above procedure using the red filter labelled
623nm. DO NOT make a movie using the red filter; record the total number of cells in the field
of view and illuminate the cells for an hour. After the light treatment record the number of
incipient and complete systrophe. Be sure to adjust your power to get the desired photon flux
of 200 μmol m<sup>-2</sup> s<sup>-1</sup> and ensure your microscope is set up for Kohler illumination.</li>
</ol>
</ol>

<h1> Questions/Write-up </h1>

<p>Answer the following questions in your lab notebook.</p>
<ol>
<li>An additional experiment was performed [8] by illuminating only half the cell at a time. The
results of this experiment are shown below:
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig11.png|700px|center]]
<b>Figure 11:</b> Half Cell Illuminations: This particular cell was illuminated under broad band-pass
irradiation with a photon flux of <b>1500μmol m<sup>-2</sup> s<sup>-1</sup>.</b> After an hour of irradiation, chloroplasts have
vacated the illuminated area.
<br clear=right>
</p>
</td></table>
<p>What can you conclude about these results? Is chloroplast translocation a protective
mechanism for the cell or rather an avoidance mechanism? Explain in a few sentences.</p></li>

<li>Imagine you had a source of UV light available and you wished to perform an additional
experiment by illuminating the cells with UV light as you did earlier with the blue and red bandpass
filters. Assume your UV light source emits at a peak wavelength of 250nm ± 10nm. Taking
into account the absorption spectra of chlorophyll shown in Figure 1 and that DNA absorbs light
at 260nm what would you expect to happen?</li>

<li>During this lab we examined the wavelength dependence of light on systrophe, however other
experiments [8] demonstrated there is also an intensity dependence as shown in the Figure
below:</li>

<table width=500 align=center><td>
<p align=center>[[File:Ct_fig13.png|500px|center]]
<b>Figure 13:</b> Intensity Dependence of Light on Incipient and Complete Systrophe.
<br clear=right>
</p>
</td></table>

<p>At a photon flux of around <b>2000μmol m<sup>-2</sup> s<sup>-1</sup></b> there is a dramatic increase in the number of
incipient systrophe. Remember incipient means the cells are in the process of undergoing
chloroplast translocation. The photon flux we receive from the sun is around <b>2000μmol m<sup>-2</sup> s<sup>-1</sup></b>.
What does this imply?</p></li>

<li>Photosynthetic organisms use wavelengths between 400nm and 800nm for photosynthesis.
From Figure 1, it can be seen that chlorophyll strongly absorbs blue and red light. Taking this
into account and your answers to questions 1 and 2 what role do you think systrophe plays in
photosynthesis? Are they related? Please discuss in a few sentences providing insights into
your thinking process.</li>
</ol>

<p>For the formal write-up for this experiment, briefly describe the purpose of the experiment, the
experimental set-up and the physics behind the biological process we’re observing, this will be your
introduction. You should type up your results into a table comparing the illumination trials using red
and blue filters. Provide a brief discussion interpreting your results. In addition you need to provide
answers to the following questions.</p>
<ol>
<li>The effect of systrophe is thought to be a defence mechanism for the cell by decreasing the
absorptive cross sectional area of chloroplasts within the cell. We can test this hypothesis by
using a simple mathematical model to represent the cell. Assume each cell of E. viridis in its
resting state is a sphere with a shell of chloroplasts along the periphery of the cell. In reality
chloroplasts are also distributed radially along the cytoplasmic strands within the cell (analogous
to the spokes of a bicycle wheel); however for simplicity we will only consider the outer shell of
chloroplasts. After the cell undergoes systrophe, the chloroplasts surround the nucleus in a
thicker shell. In the table below some data are provided of cellular dimensions for 3 arbitrary
cells before and after systrophe using time lapsed images <ref>Web-published: http://www.yorku.ca/planters/student_reports/viridis_systrophe.pdf (accessed
9 September 2010).</ref>.

<table width=500 align=center>
<tr><td><p align=center><b>Table 1:</b> Cellular Dimensions before and after Systrophe.</p></td></tr>

<tr><td><table width=500 align=center border>
<tr><td></td><td>Cell 1</td><td>Cell 2</td><td>Cell 3</td></tr>
<tr><td>Cell Diameter (μm)</td><td>122.5</td><td>108.5</td><td>130.5</td></tr>
<tr><td>Systrophe Diameter (μm)</td><td>80.1</td><td>64.7</td><td>103.8</td></tr>
<tr><td>Nucleus Diameter (μm)</td><td>41.8</td><td>45.8</td><td>61.3</td></tr>
<tr><td>Thickness of Chloroplast Shell
(Before Systrophe) (μm)</td><td>4.6</td><td>5.5</td><td>5.3</td></tr>
</table>

</td></tr></table>

<p>Use the data in Table 1 as well as the physical properties of the cells included in the
introductions and the Beer-Lambert Law (Equation 1.3) to fill in the table below:</p>
<table width=150 align=center><td>
<p align=justify>[[File:Ct_eqn1_3.png|150px|center]]
<br clear=right>
</p>
</td></table>
<p>Where I is the transmitted intensity of light passing through the cell, Io is the initial intensity your
light source, ε is the extinction coefficient of chlorophyll, C is the concentration of chlorophyll
and l is path length which the light passes through.</p>
<table width=700 align=center><td>
<p align=center><b>Table 2: </b> Absorptive Properties of Cells before and after Systrophe.
[[File:Ct_tab2.png|650px|center]]
<br clear=right>
</p>
</td></table>

<p>Based on your results in Table 2, does systrophe provide a protective mechanism for the cell by
decreasing the absorptive cross sectional area? Why would an increase in absorption
potentially be damaging for the cell? Consider both absorption and cross sectional area.</p>
</li>

<li>Open the spreadsheet titled <b>“Viridis_Systrophe_Spectrum.xls”</b>; this is an emission spectra of
the tungsten halogen bulb used for illuminating the cell. Calculate the total energy the cells
would receive from the whole spectrum of light emitted from this source by normalization. You
can use the spreadsheet to do your calculations and may wish to attach it to your lab report.</li>
<li>Below is the emission spectrum from the tungsten halogen bulb used for illuminations. From
the Planck Blackbody Distribution Law derive Wien’s Displacement Law which relates the colour
temperature of a blackbody to the peak wavelength it emits at. If we treat tungsten as a
blackbody, what is the color temperature associated with the peak wavelength observed in
Figure 12? Does this make sense considering the melting point of tungsten is about 3350K?
What correction do we need to take into account when deriving Wien’s Displacement Law from
Planck’s Blackbody Distribution Law to account for the unusual high melting point implied from
Figure 12?</li>

<table width=500 align=center><td>
<p align=center>[[File:Ct_fig12.png|500px|center]]
<b>Figure 12:</b> Tungsten Halogen Emission Spectrum.
<br clear=right>
</p>
</td></table>

<li>During systrophe chloroplasts are translated across the cell by molecular motors. It isn’t known
exactly which molecular motors are responsible for chloroplast translocation, however it should
be noted chloroplast translocation can be inhibited when cells of <i>E. viridis</i> are treated with 2,4 –
Dinitrophenol (DNP) [4]. The structure of DNP is shown in Figure 14.

<table width=400 align=center><td>
<p align=center>[[File:Ct_fig14.png|200px|center]]
<b>Figure 14:</b> Structure of 2,4 Dinitrophenol.
<br clear=right>
</p>
</td></table>
DNP acts as an uncoupling agent by disrupting the proton gradient the cell uses to create ATP.
Any energy the cell would have gained from ATP production is lost as heat. ATP provides the
energy for molecular motors to shuttle organelles across the cell. Based on the structure of
DNP, taking into account hydrophilicity and hydrophobicity and any partial charges on the
molecule explain how DNP inhibits chloroplast translocation. Does this provide any insight into
the energy mechanisms involved during systrophe?
</li>

<h1>References</h1>
<References/>

File:Chlorophyll a spectrum.png

2012-08-10T11:34:55Z

Rlew: Chlorophyll a spectrum in a methanol solvent.

Chlorophyll a spectrum in a methanol solvent.

Main Page/BPHS 4090/Choloplast Translocation

2012-08-10T11:34:21Z

Rlew:

<h1>Required Components</h1>
<ol>
<li>[[Media:M3_Viridis.JPG|Stock culture of <i>Eremosphaera viridis</i>]]</li>
<li>[[Media:Slide_Tools.JPG|Glass slides, cover slips and micropipette]]</li>
<li>Nikon Optiphot Microscope</li>
<li>Radiometric Probe (Model S471 Portable Optometer, UDT Instruments)</li>
<li>[[Media:CCD.JPG|Cool Snap CCD Camera and MicroManager software]]</li>
<li>[[Media:M3_Band_Pass.JPG|Blue and red band-pass filters]]</li>
</ol>

<h1>Objective</h1>
<p>To observe the wavelength dependence of light on chloroplast translocation in the acidophile green
algae <i>Eremosphaera viridis</i> as well as describing a mechanism behind this response.</p>

<h1>Introduction</h1>
<p>Electromagnetic radiation is the driving force for photosynthesis. Plants and other autotrophs convert
the energy of light into chemical energy used for synthesizing carbohydrates and other organic
compounds, which heterotrophic organisms use for energy and other nutritional requirements.
Photosynthesis is a unique process performed by plants, algae, and some species of bacteria. It consists
of a series of oxidation and reduction reactions; the basic chemical formula is shown below:
<table width=380 align=center><td>
<p align=justify>[[File:Ct_eqn0.png|340px|center]]
<br clear=right>
</p>
</td></table>
Photosynthesis would not be possible without the energy derived from light. Electromagnetic radiation
enters the earth’s atmosphere from the sun where it is harvested by photosynthetic organisms.
Wavelengths between 400 and 800 nm are used by photosynthetic organisms. The sun can be
approximated as a near perfect blackbody with a surface temperature of 5800 K described by Planck’s
Black Body Radiation Law, shown in Equation 1.1
<table width=300 align=center><td>
<p align=justify>[[File:Ct_eqn1_1.png|280px|right]]
<br clear=right>
</p>
</td></table>
Where U (λ, T) has units of J m<sup>-2</sup> s<sup>-1</sup>, h is Planck’s constant, K is Boltzmann’s Constant and c is the speed
of light. The color temperature of the sun corresponds to a peak wavelength emission of about 500nm
according to Wien’s Displacement Law (Equation 1.2)
<table width=180 align=center><td>
<p align=justify>[[File:Ct_eqn1_2.png|160px|right]]
<br clear=right>
</p>
</td></table>
Where b = 2.898 x 10<sup>-3</sup> in units of m*K.</p>

<p>The first step in the conversion of the energy of photons to chemical energy is absorption by
photosynthetic pigments, principally chlorophyll. The absorption spectrum of chlorophyll a is shown in
Figure 1 <ref>http://www.bio.davidson.edu/courses/Bio111/Bio111LabMan/lab1fig3.gif</ref>.</p>

<table width=800 align=center><td>
<p align=justify>[[File:Chlorophyll_a_spectrum.png|300px|left]]
<b>Absorption Spectrum of Chlorophyll a:</b> Chlorophyll a strongly absorbs
light at 465nm and 665nm. There is minimal
absorption of UV light as well as green/yellow light
which appears in the range of 500nm-600nm. These
absorbances are for chlorophyll isolated in a
solvent of a well-defined dielectric. In the cell,
additional pigments and heterogeneous dielectric
causes much broader peaks.
<br clear=right>
</p>
</td></table>

<p><b>DETERMINING EXCITATION AND EMISSION SPECTRA</b><br>
<table align=right border=0 width=400>
<tr><td>[[File:Fluorescence_scans.png|200px|left]]</td>
<td>Here is an example of absorbance/fluorescence scans of whole algal cells and acetone/water extracted chlorophyll to give you an idea about the kind of absorbance that occurs
during light irradiation of your cells.</td></tr>
</table>

<p>Chlorophyll is located inside the thylakoid vesicles in the inner membrane of the chloroplast. After
absorbing a photon, an electron in chlorophyll transitions to an excited state. The excited state electron
(exciton) is transferred to a photosynthetic reaction center to begin the flow of electrons through the
electron transport chain, eventually to produce ATP (from a transmembrane H<sup>+</sup> gradient) and reducing
equivalents (NADP + 2e<sup>–</sup> + 2H<sup>+</sup> —> NADPH + H<sup>+</sup>). Chlorophyll, now with one less electron, is chemically
unstable and receives an electron from a water molecule (H<sub>2</sub>O). With the loss of 4 electrons from two
molecules of water, molecular oxygen (O<sub>2</sub>) is produced, as well as 4H<sup>+</sup> used for ATP synthesis. Oxygen,
the waste product of photosynthesis, is vital for heterotrophs in the process of cellular respiration.</p>

<p>As light intensity increases, so do absorption events and the generation of excited state chlorophylls,
and downstream electron transport. At a high enough light intensity, these can cause the formation of a
variety of undesirable oxidative products that can damage the photosynthetic apparatus. Examples
include triplet state chlorophyll, and various reactive oxygen species (through direct reduction of O<sub>2</sub> to
O<sub>2</sub><sup>–</sup>, and subsequent formation of H<sub>2</sub>O<sub>2</sub>). The general term photo-oxidation is used to describe this
damaging process. There are protective mechanisms to avoid oxidative damage <ref>Li, Z., Wakao, S., Fischer, B.B., Niyogi, K.K. 2009. Sensing and responding to excess light. Annual Review of Plant Biology <b>60</b>: 239–260.</ref>; one of these may be
to decrease the absorptive cross-sectional area of chloroplasts by changing their location in the cell <ref>Kasahara M., Kagawa, T., Oikawa, K., Suetsugu, N., Miyao, M., Wada, M. 2002. Chloroplast avoidance movement reduces photodamage in plants. Nature 420: 829–832</ref>. The phenomenon known as systrophe is described as the accumulation of cytoplasmic organelles
around the nucleus <ref>Weidinger, M. 1980. The inhibition of systrophe by cytochalasin B. Protoplasma <b>102</b>: 167–170.</ref><ref>Weidinger, M. 1982. The inhibition of systrophe in different organisms. Protoplasma <b>110</b>: 71–
74.</ref><ref>Weidinger, M., Ruppel, H.G. 1985. Ca<sup>2+</sup> requirement for a blue-light-induced chloroplast
translocation in <i>Eremosphaera viridis</i>. Protoplasma <b>124</b>: 184–187.</ref>. Systrophe of the chloroplasts is observed in the unicellular green algae <i>Eremosphaera viridis</i> when the cell is exposed to intensities of light greater than it would experience in
its natural environment.</p>
<p>During this experiment we will be examining chloroplast translocation in the unicellular green algae
<i>Eremosphaera viridis</i>, in particular we will be looking at the wavelength dependence of light on
systrophe. A typical cell of <i>E. viridis</i> is shown in Figure 2 during the course of a light treatment protocol.</p>

<table width=800 align=center><td>
<p align=justify>[[File:Ct_fig2.png|780px|center]]
<b>Figure 2: Examples of incipient and Complete Systrophe in <i>Eremosphaera
viridis</i></b> This particular cell was illuminated with a photon flux of 200 μmol m<sup>-2</sup> s<sup>-1</sup> using a blue bandpass filter with a peak wavelength of 441nm ± 10nm.
<br clear=right>
</p>
</td></table>

<p>From Figure 2, it can be seen that the systrophe effect is quite dramatic! At 0 minutes is how the cell
would appear in its resting state, at 20 minutes into the light treatment protocol there are signs of
chloroplast translocation, this cell would be classified as incipient systrophe since it is in the process of
undergoing systrophe. Lastly at 60 minutes into the light treatment the cell would be classified as
complete systrophe. Notice at 60 minutes the cytoplasmic strands which chloroplasts migrate along
during light treatments are clearly visible!</p>
<p>An <i>Eremosphaera viridis</i> cell has about 400 chloroplasts, based on counting of medial sections of
fluorescence images. In a normal chloroplast, there is about 9•10<sup>–13</sup> g of chlorophyll and about 6.7 • 10<sup>8</sup>
chlorophyll molecules per chloroplast. The molecular weight of chlorophyll a is about 894; its extinction
coefficient is 1.2 • 10<sup>5</sup> M<sup>–1</sup> cm<sup>–1</sup> at 430nm <ref>Lawlor, D.W. 2001. Photosynthesis. Third edition. Springer-Verlag, New York. Chapters 3 and
4.</ref>.</p>

<h1>Materials and Methods </h1>

<p>The procedure is divided into four sections:
<ol style="list-style-type:upper-latin">
<li>Preparing a sample slide</li>
<li>Microscope Set-Up and Kohler Illumination</li>
<li>Light Treatments</li>
<li>Image Processing</li>
</ol>
Please follow them in order; each section has its own set of step by step instructions.</p>

<ol style="list-style-type:upper-latin">
<h2><li>Preparing a Sample Slide</h2>
<ol>
<li>Take a new glass slide and trace the outline of the cover slip using a permanent black marker as
shown in Figure 3A.</li>
<li>Spread a thin layer of Vaseline as shown in Figure 3C & D along the outline you just traced using
the syringe (Figure 3B), this will prevent your cell sample from drying out under high light
tensities</li>
<li>Aliquot 10μL of <i>E. viridis</i> in the outline you just traced using a micropipette as shown in Figure
3E</li>
<li>Gently place the cover slip on top of the Vaseline layer, being careful not to crush your cell
sample</li>
<li>Now the slide is ready to be used for light treatments (Figure 3F).</li>
<table width=600 align=center><td>
<p align=justify>[[File:Ct_fig3.png|580px|center]]
<b>Figure 3</b>
<br clear=right>
</p>
</td></table>
</ol>
</li>
<h2><li>Microscope Set-up and Kohler Illumination:</h2>
<ol>
<li>This experiment will be performed using bright-field microscopy. Ensure the ring on the
condenser diaphragm is set at 0 (not PH 1 or PH 2), because the phase ring at the PH1 or PH2
setting attenuates the light.</li>
<li>Once your specimen slide is prepared, bring it to the microscope and focus on the specimen
under the x10 objective.</li>
<li>Next close the field diaphragm all the way shut so you can see the edges, they may appear
blurry.</li>
<li>Use the condenser focus knob to bring the edges of the field diaphragm into the best possible
focus.</li>
<li>Next use the condenser-centering screws to bring the closed field diaphragm into the center of
the field of view.</li>
<li>Then open the field diaphragm such that it is slightly larger than the field of view.</li>
<li>You are now set up for Kohler illumination.</li>
</ol>
</li>
<h2><li>Light Treatments</h2>
<p>The wavelength dependence of light on chloroplast translocation will be investigated. From the
absorption spectrum of chlorophyll shown in Figure 1, it is known the chlorophyll strongly absorbs both
blue and red light, thus for the wavelength experiments we will be using blue and red filters band-pass
filters, their transmissions are shown in Figure 4. The blue band-pass filter emits light at a peak
wavelength of 441nm ± 10nm. The red filter is a band-pass filter with a tail at longer wavelengths. This
filter has a peak wavelength of 623nm ± 45nm.</p>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig4.png|450px|left]]
<b>Figure 4: Emission Spectra of Band-pass Filters:</b>The blue bandpass
filter was fit to a
Gaussian function and the
red band-pass filter was fit
to a Lorentzian Function to
account for the long tail
end. Both fit functions are
shown in black.
<br clear=right>
</p>
</td></table>

<p>The cells will be illuminated with both blue and red light to determine the wavelength dependence of
light on chloroplast translocation as described in the following outline:</p>
<ol>
<li>Take the blue band-pass filter labelled 441nm and place it above the light source under the
condenser diaphragm.</li>
<li>Double click on the Desktop folder called <b>BPHS_4090_Viridis_Systrophe</b>, and then open the
Excel spreadsheet called <b>“Photon_Flux_Calculator.xls”</b>. This spreadsheet is already
programmed to do all the difficult calculations for you to save you time in the lab!</li>
<li>With the blue filter in place turn on the radiometer, use the probe to measure the output power
from the light source; this is done by removing the sample slide and placing the probe in the
light path where the objective is. You may have to remove one of the eyepieces to get the
probe in the light path, do this carefully and ensure you leave the x10 objective (labelled Fluor
10) in place, since this will be used for the light treatments. Ensure the probe is collecting all the
light. You should measure a power reading in microwatts; enter this into the spreadsheet to get
a photon flux as an output in units of μmol m<sup>-2</sup> s<sup>-1</sup> as shown in Figure 5.</li>

<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig5.png|650px|left]]
<b>Figure 5:</b> The flux calculator has separate sections for the red and blue bandpass
filters. The cells will feel different energies corresponding to the different wavelengths of light
according to the Planck relationship <i>E = hc/λ </i>.
<br clear=right>
</p>
</td></table>

<li>For the wavelength experiment a photon flux of around <b> 200μmol m<sup>-2</sup> s<sup>-1</sup></b> is ideal to observe the
effect of chloroplast translocation. Adjust the voltage dial knob on the microscope until you get
a power reading which gives you a photon flux close to this value,<b>once you do so, record your
input power.</b></li>
<li>Next place the cell sample back in the field of view, drop the neutral density filter labelled ND 2
in the light path to avoid pre-treating the cells with light. You’ll have to adjust the microscope
again for Kohler Illumination, but it shouldn’t take much adjusting at this point.</li>
<li>Move the microscope stage to capture as large of a cell population as possible, try to aim for 20
or more cells if possible by looking through the eyepiece as you move the stage.</li>
<li>Ensure your cell sample is in good focus, you’re now ready to make a movie of chloroplast
translocation.</li>
</ol>

<h2><li>Making a Movie of Chloroplast Translocation:</h2>
<ol>
<li>Turn on the CoolSnap CCD camera.</li>
<li>Open MicroManager 1.3 by double clicking the icon on the desktop, you’ll notice the program
ImageJ (Figure 6) opens as well, this will be used later for image processing.
Figure 6 – MicroManager Software</li>
<table width=600 align=center><td>
<p align=justify>[[File:Ct_fig6.png|550px|left]]
<b>Figure 6:</b> MicroManager Software.
<br clear=right>
</p>
</td></table>
<li>Click on Live Feed and turn the eyepiece mounting block about 45 degrees to the left, this allows
light to pass through to the CCD camera and will project a live image onto the computer
monitor.</li>
<li>Next lift up the ND2 filter and adjust the exposure time, start by setting it to 1ms, you’ll also
notice the cells on the screen are out of focus, use the fine focus adjustment until you get crisp
edges along the periphery of the cell.</li>
<li>Adjust the exposure until you get a live image that isn’t oversaturated (Figure 7).</li>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig7.png|650px|left]]
<b>Figure 7:</b> MicroManager Live Feed.
<br clear=right>
</p>
</td></table>
<li>Due to the limited field of view, you’ll only be able to get about 1 cell in the live feed. Move the
microscope stage slowly (since the response from the CCD is slightly delayed) until you get 1 cell
in the field of view as shown in Figure 7.</li>
<li>Click on “Multi-D acq.” Then set the number to 360 and interval to 10 seconds, and then hit
acquire (Figure 8). This tells the program to take an image every 10 seconds; this will eventually
give you a movie 60 minutes long, which should allow you to see some significant chloroplast
translocation as shown in Figure 2.</li>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig8.png|450px|center]]
<b>Figure 8:</b> Multi-Dimensional Acquisition.
<br clear=right>
</p>
</td></table>
<li>Once finished click on the save icon. Save your data in the directory
<b>C:\BPHS4090_Viridis_Systrophe, label your data by your name, date and title of work.</b> This
will create a file folder with all the images you acquired.</li>
<table width=300 align=center><td>
<p align=justify>[[File:Ct_fig9.png|250px|center]]
<b>Figure 9:</b> Sequence Options.
<br clear=right>
</p>
</td></table>
<li>Next you’ll be using ImageJ. Click “File->Import->Image
Sequence”, double click on your file folder and select
one of the images and press Open. Another window
will open called “Sequence Options” (Figure 9), it will
verify the number of images you have and open them
as a stack, make sure the box titled “Sort Names
Numerically” is checked and click Ok.</li>
<li>This may take a little while, once all the images are
loaded, use the mouse cursor icon to make a box
around your cell and click on “Image->Crop”. Every
image you imported will be cropped.</li>
<li>Next click on “Image->Adjust->Brightness/Contrast”,
another window called “B&C” will open; adjust the
settings to ensure your images look the best they can.
Once happy with the results click on Set, a new window
will open, make sure the “Propagate to all open
images” box is checked and click OK (Figure 10). This
will adjust the brightness and contrast for every image
in your imported stack.</li>

<table width=400 align=center><td>
<p align=justify>[[File:Ct_fig10.png|350px|center]]
<b>Figure 10:</b> Adjusting Image Brightness and Contrast.
<br clear=right>
</p>
</td></table>
<li>Once your image is adjusted for proper brightness and contrast select “File->Save as->AVI”,
select a frame rate of about 10fps, this will make a movie 36 seconds long assuming you took
360 frames. A movie at this speed will dramatically show chloroplast translocation assuming the
one cell you chose to view undergoes the effect. Save the AVI in the directory
<b>C:\BPHS4090_Viridis_Systrophe by your name, date and title of work.</b></li>
<li><b>Copy your movie and the file called “Viridis_Systrophe_Spectrum.xls” onto a USB Key, this
spreadsheet will be used for one of the questions. You can find the file in the desktop folder
called “BPHS_4090_Viridis_Systrophe”.</b></li>
<li>Look at your cell sample through the field of view and record how many cells are incipient
systrophe and how many are complete systrophe using the criteria in Figure 2.</li>
<li>Once this is done discard your cell sample slide in the bin labelled “Sharps”</li>
<li>Next prepare a new sample slide and repeat the above procedure using the red filter labelled
623nm. DO NOT make a movie using the red filter; record the total number of cells in the field
of view and illuminate the cells for an hour. After the light treatment record the number of
incipient and complete systrophe. Be sure to adjust your power to get the desired photon flux
of 200 μmol m<sup>-2</sup> s<sup>-1</sup> and ensure your microscope is set up for Kohler illumination.</li>
</ol>
</ol>

<h1> Questions/Write-up </h1>

<p>Answer the following questions in your lab notebook.</p>
<ol>
<li>An additional experiment was performed [8] by illuminating only half the cell at a time. The
results of this experiment are shown below:
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig11.png|700px|center]]
<b>Figure 11:</b> Half Cell Illuminations: This particular cell was illuminated under broad band-pass
irradiation with a photon flux of <b>1500μmol m<sup>-2</sup> s<sup>-1</sup>.</b> After an hour of irradiation, chloroplasts have
vacated the illuminated area.
<br clear=right>
</p>
</td></table>
<p>What can you conclude about these results? Is chloroplast translocation a protective
mechanism for the cell or rather an avoidance mechanism? Explain in a few sentences.</p></li>

<li>Imagine you had a source of UV light available and you wished to perform an additional
experiment by illuminating the cells with UV light as you did earlier with the blue and red bandpass
filters. Assume your UV light source emits at a peak wavelength of 250nm ± 10nm. Taking
into account the absorption spectra of chlorophyll shown in Figure 1 and that DNA absorbs light
at 260nm what would you expect to happen?</li>

<li>During this lab we examined the wavelength dependence of light on systrophe, however other
experiments [8] demonstrated there is also an intensity dependence as shown in the Figure
below:</li>

<table width=500 align=center><td>
<p align=center>[[File:Ct_fig13.png|500px|center]]
<b>Figure 13:</b> Intensity Dependence of Light on Incipient and Complete Systrophe.
<br clear=right>
</p>
</td></table>

<p>At a photon flux of around <b>2000μmol m<sup>-2</sup> s<sup>-1</sup></b> there is a dramatic increase in the number of
incipient systrophe. Remember incipient means the cells are in the process of undergoing
chloroplast translocation. The photon flux we receive from the sun is around <b>2000μmol m<sup>-2</sup> s<sup>-1</sup></b>.
What does this imply?</p></li>

<li>Photosynthetic organisms use wavelengths between 400nm and 800nm for photosynthesis.
From Figure 1, it can be seen that chlorophyll strongly absorbs blue and red light. Taking this
into account and your answers to questions 1 and 2 what role do you think systrophe plays in
photosynthesis? Are they related? Please discuss in a few sentences providing insights into
your thinking process.</li>
</ol>

<p>For the formal write-up for this experiment, briefly describe the purpose of the experiment, the
experimental set-up and the physics behind the biological process we’re observing, this will be your
introduction. You should type up your results into a table comparing the illumination trials using red
and blue filters. Provide a brief discussion interpreting your results. In addition you need to provide
answers to the following questions.</p>
<ol>
<li>The effect of systrophe is thought to be a defence mechanism for the cell by decreasing the
absorptive cross sectional area of chloroplasts within the cell. We can test this hypothesis by
using a simple mathematical model to represent the cell. Assume each cell of E. viridis in its
resting state is a sphere with a shell of chloroplasts along the periphery of the cell. In reality
chloroplasts are also distributed radially along the cytoplasmic strands within the cell (analogous
to the spokes of a bicycle wheel); however for simplicity we will only consider the outer shell of
chloroplasts. After the cell undergoes systrophe, the chloroplasts surround the nucleus in a
thicker shell. In the table below some data are provided of cellular dimensions for 3 arbitrary
cells before and after systrophe using time lapsed images <ref>Web-published: http://www.yorku.ca/planters/student_reports/viridis_systrophe.pdf (accessed
9 September 2010).</ref>.

<table width=500 align=center>
<tr><td><p align=center><b>Table 1:</b> Cellular Dimensions before and after Systrophe.</p></td></tr>

<tr><td><table width=500 align=center border>
<tr><td></td><td>Cell 1</td><td>Cell 2</td><td>Cell 3</td></tr>
<tr><td>Cell Diameter (μm)</td><td>122.5</td><td>108.5</td><td>130.5</td></tr>
<tr><td>Systrophe Diameter (μm)</td><td>80.1</td><td>64.7</td><td>103.8</td></tr>
<tr><td>Nucleus Diameter (μm)</td><td>41.8</td><td>45.8</td><td>61.3</td></tr>
<tr><td>Thickness of Chloroplast Shell
(Before Systrophe) (μm)</td><td>4.6</td><td>5.5</td><td>5.3</td></tr>
</table>

</td></tr></table>

<p>Use the data in Table 1 as well as the physical properties of the cells included in the
introductions and the Beer-Lambert Law (Equation 1.3) to fill in the table below:</p>
<table width=150 align=center><td>
<p align=justify>[[File:Ct_eqn1_3.png|150px|center]]
<br clear=right>
</p>
</td></table>
<p>Where I is the transmitted intensity of light passing through the cell, Io is the initial intensity your
light source, ε is the extinction coefficient of chlorophyll, C is the concentration of chlorophyll
and l is path length which the light passes through.</p>
<table width=700 align=center><td>
<p align=center><b>Table 2: </b> Absorptive Properties of Cells before and after Systrophe.
[[File:Ct_tab2.png|650px|center]]
<br clear=right>
</p>
</td></table>

<p>Based on your results in Table 2, does systrophe provide a protective mechanism for the cell by
decreasing the absorptive cross sectional area? Why would an increase in absorption
potentially be damaging for the cell? Consider both absorption and cross sectional area.</p>
</li>

<li>Open the spreadsheet titled <b>“Viridis_Systrophe_Spectrum.xls”</b>; this is an emission spectra of
the tungsten halogen bulb used for illuminating the cell. Calculate the total energy the cells
would receive from the whole spectrum of light emitted from this source by normalization. You
can use the spreadsheet to do your calculations and may wish to attach it to your lab report.</li>
<li>Below is the emission spectrum from the tungsten halogen bulb used for illuminations. From
the Planck Blackbody Distribution Law derive Wien’s Displacement Law which relates the colour
temperature of a blackbody to the peak wavelength it emits at. If we treat tungsten as a
blackbody, what is the color temperature associated with the peak wavelength observed in
Figure 12? Does this make sense considering the melting point of tungsten is about 3350K?
What correction do we need to take into account when deriving Wien’s Displacement Law from
Planck’s Blackbody Distribution Law to account for the unusual high melting point implied from
Figure 12?</li>

<table width=500 align=center><td>
<p align=center>[[File:Ct_fig12.png|500px|center]]
<b>Figure 12:</b> Tungsten Halogen Emission Spectrum.
<br clear=right>
</p>
</td></table>

<li>During systrophe chloroplasts are translated across the cell by molecular motors. It isn’t known
exactly which molecular motors are responsible for chloroplast translocation, however it should
be noted chloroplast translocation can be inhibited when cells of <i>E. viridis</i> are treated with 2,4 –
Dinitrophenol (DNP) [4]. The structure of DNP is shown in Figure 14.

<table width=400 align=center><td>
<p align=center>[[File:Ct_fig14.png|200px|center]]
<b>Figure 14:</b> Structure of 2,4 Dinitrophenol.
<br clear=right>
</p>
</td></table>
DNP acts as an uncoupling agent by disrupting the proton gradient the cell uses to create ATP.
Any energy the cell would have gained from ATP production is lost as heat. ATP provides the
energy for molecular motors to shuttle organelles across the cell. Based on the structure of
DNP, taking into account hydrophilicity and hydrophobicity and any partial charges on the
molecule explain how DNP inhibits chloroplast translocation. Does this provide any insight into
the energy mechanisms involved during systrophe?
</li>

<h1>References</h1>
<References/>

Main Page/BPHS 4090/Choloplast Translocation

2012-08-10T11:32:23Z

Rlew:

<h1>Required Components</h1>
<ol>
<li>[[Media:M3_Viridis.JPG|Stock culture of <i>Eremosphaera viridis</i>]]</li>
<li>[[Media:Slide_Tools.JPG|Glass slides, cover slips and micropipette]]</li>
<li>Nikon Optiphot Microscope</li>
<li>Radiometric Probe (Model S471 Portable Optometer, UDT Instruments)</li>
<li>[[Media:CCD.JPG|Cool Snap CCD Camera and MicroManager software]]</li>
<li>[[Media:M3_Band_Pass.JPG|Blue and red band-pass filters]]</li>
</ol>

<h1>Objective</h1>
<p>To observe the wavelength dependence of light on chloroplast translocation in the acidophile green
algae <i>Eremosphaera viridis</i> as well as describing a mechanism behind this response.</p>

<h1>Introduction</h1>
<p>Electromagnetic radiation is the driving force for photosynthesis. Plants and other autotrophs convert
the energy of light into chemical energy used for synthesizing carbohydrates and other organic
compounds, which heterotrophic organisms use for energy and other nutritional requirements.
Photosynthesis is a unique process performed by plants, algae, and some species of bacteria. It consists
of a series of oxidation and reduction reactions; the basic chemical formula is shown below:
<table width=380 align=center><td>
<p align=justify>[[File:Ct_eqn0.png|340px|center]]
<br clear=right>
</p>
</td></table>
Photosynthesis would not be possible without the energy derived from light. Electromagnetic radiation
enters the earth’s atmosphere from the sun where it is harvested by photosynthetic organisms.
Wavelengths between 400 and 800 nm are used by photosynthetic organisms. The sun can be
approximated as a near perfect blackbody with a surface temperature of 5800 K described by Planck’s
Black Body Radiation Law, shown in Equation 1.1
<table width=300 align=center><td>
<p align=justify>[[File:Ct_eqn1_1.png|280px|right]]
<br clear=right>
</p>
</td></table>
Where U (λ, T) has units of J m<sup>-2</sup> s<sup>-1</sup>, h is Planck’s constant, K is Boltzmann’s Constant and c is the speed
of light. The color temperature of the sun corresponds to a peak wavelength emission of about 500nm
according to Wien’s Displacement Law (Equation 1.2)
<table width=180 align=center><td>
<p align=justify>[[File:Ct_eqn1_2.png|160px|right]]
<br clear=right>
</p>
</td></table>
Where b = 2.898 x 10<sup>-3</sup> in units of m*K.</p>

<p>The first step in the conversion of the energy of photons to chemical energy is absorption by
photosynthetic pigments, principally chlorophyll. The absorption spectrum of chlorophyll a is shown in
Figure 1 <ref>http://www.bio.davidson.edu/courses/Bio111/Bio111LabMan/lab1fig3.gif</ref>.</p>

<table width=800 align=center><td>
<p align=justify>[[File:Ct_fig1.png|300px|left]]
<b>Absorption Spectrum of Chlorophyll a:</b> Chlorophyll a strongly absorbs
light at 465nm and 665nm. There is minimal
absorption of UV light as well as green/yellow light
which appears in the range of 500nm-600nm. These
absorbances are for chloroplasts isolated in a
solvent of a well-defined dielectric. In the cell,
additional pigments and heterogeneous dielectric
causes much broader peaks.
<br clear=right>
</p>
</td></table>

<p><b>DETERMINING EXCITATION AND EMISSION SPECTRA</b><br>
<table align=right border=0 width=400>
<tr><td>[[File:Fluorescence_scans.png|200px|left]]</td>
<td>Here is an example of fluorescence scans of whole algal cells and acetone/water extracted chlorophyll to give you an idea about the kind of absorbance that occurs
during light irradiation of your cells.</td></tr>
</table>

<p>Chlorophyll is located inside the thylakoid vesicles in the inner membrane of the chloroplast. After
absorbing a photon, an electron in chlorophyll transitions to an excited state. The excited state electron
(exciton) is transferred to a photosynthetic reaction center to begin the flow of electrons through the
electron transport chain, eventually to produce ATP (from a transmembrane H<sup>+</sup> gradient) and reducing
equivalents (NADP + 2e<sup>–</sup> + 2H<sup>+</sup> —> NADPH + H<sup>+</sup>). Chlorophyll, now with one less electron, is chemically
unstable and receives an electron from a water molecule (H<sub>2</sub>O). With the loss of 4 electrons from two
molecules of water, molecular oxygen (O<sub>2</sub>) is produced, as well as 4H<sup>+</sup> used for ATP synthesis. Oxygen,
the waste product of photosynthesis, is vital for heterotrophs in the process of cellular respiration.</p>

<p>As light intensity increases, so do absorption events and the generation of excited state chlorophylls,
and downstream electron transport. At a high enough light intensity, these can cause the formation of a
variety of undesirable oxidative products that can damage the photosynthetic apparatus. Examples
include triplet state chlorophyll, and various reactive oxygen species (through direct reduction of O<sub>2</sub> to
O<sub>2</sub><sup>–</sup>, and subsequent formation of H<sub>2</sub>O<sub>2</sub>). The general term photo-oxidation is used to describe this
damaging process. There are protective mechanisms to avoid oxidative damage <ref>Li, Z., Wakao, S., Fischer, B.B., Niyogi, K.K. 2009. Sensing and responding to excess light. Annual Review of Plant Biology <b>60</b>: 239–260.</ref>; one of these may be
to decrease the absorptive cross-sectional area of chloroplasts by changing their location in the cell <ref>Kasahara M., Kagawa, T., Oikawa, K., Suetsugu, N., Miyao, M., Wada, M. 2002. Chloroplast avoidance movement reduces photodamage in plants. Nature 420: 829–832</ref>. The phenomenon known as systrophe is described as the accumulation of cytoplasmic organelles
around the nucleus <ref>Weidinger, M. 1980. The inhibition of systrophe by cytochalasin B. Protoplasma <b>102</b>: 167–170.</ref><ref>Weidinger, M. 1982. The inhibition of systrophe in different organisms. Protoplasma <b>110</b>: 71–
74.</ref><ref>Weidinger, M., Ruppel, H.G. 1985. Ca<sup>2+</sup> requirement for a blue-light-induced chloroplast
translocation in <i>Eremosphaera viridis</i>. Protoplasma <b>124</b>: 184–187.</ref>. Systrophe of the chloroplasts is observed in the unicellular green algae <i>Eremosphaera viridis</i> when the cell is exposed to intensities of light greater than it would experience in
its natural environment.</p>
<p>During this experiment we will be examining chloroplast translocation in the unicellular green algae
<i>Eremosphaera viridis</i>, in particular we will be looking at the wavelength dependence of light on
systrophe. A typical cell of <i>E. viridis</i> is shown in Figure 2 during the course of a light treatment protocol.</p>

<table width=800 align=center><td>
<p align=justify>[[File:Ct_fig2.png|780px|center]]
<b>Figure 2: Examples of incipient and Complete Systrophe in <i>Eremosphaera
viridis</i></b> This particular cell was illuminated with a photon flux of 200 μmol m<sup>-2</sup> s<sup>-1</sup> using a blue bandpass filter with a peak wavelength of 441nm ± 10nm.
<br clear=right>
</p>
</td></table>

<p>From Figure 2, it can be seen that the systrophe effect is quite dramatic! At 0 minutes is how the cell
would appear in its resting state, at 20 minutes into the light treatment protocol there are signs of
chloroplast translocation, this cell would be classified as incipient systrophe since it is in the process of
undergoing systrophe. Lastly at 60 minutes into the light treatment the cell would be classified as
complete systrophe. Notice at 60 minutes the cytoplasmic strands which chloroplasts migrate along
during light treatments are clearly visible!</p>
<p>An <i>Eremosphaera viridis</i> cell has about 400 chloroplasts, based on counting of medial sections of
fluorescence images. In a normal chloroplast, there is about 9•10<sup>–13</sup> g of chlorophyll and about 6.7 • 10<sup>8</sup>
chlorophyll molecules per chloroplast. The molecular weight of chlorophyll a is about 894; its extinction
coefficient is 1.2 • 10<sup>5</sup> M<sup>–1</sup> cm<sup>–1</sup> at 430nm <ref>Lawlor, D.W. 2001. Photosynthesis. Third edition. Springer-Verlag, New York. Chapters 3 and
4.</ref>.</p>

<h1>Materials and Methods </h1>

<p>The procedure is divided into four sections:
<ol style="list-style-type:upper-latin">
<li>Preparing a sample slide</li>
<li>Microscope Set-Up and Kohler Illumination</li>
<li>Light Treatments</li>
<li>Image Processing</li>
</ol>
Please follow them in order; each section has its own set of step by step instructions.</p>

<ol style="list-style-type:upper-latin">
<h2><li>Preparing a Sample Slide</h2>
<ol>
<li>Take a new glass slide and trace the outline of the cover slip using a permanent black marker as
shown in Figure 3A.</li>
<li>Spread a thin layer of Vaseline as shown in Figure 3C & D along the outline you just traced using
the syringe (Figure 3B), this will prevent your cell sample from drying out under high light
tensities</li>
<li>Aliquot 10μL of <i>E. viridis</i> in the outline you just traced using a micropipette as shown in Figure
3E</li>
<li>Gently place the cover slip on top of the Vaseline layer, being careful not to crush your cell
sample</li>
<li>Now the slide is ready to be used for light treatments (Figure 3F).</li>
<table width=600 align=center><td>
<p align=justify>[[File:Ct_fig3.png|580px|center]]
<b>Figure 3</b>
<br clear=right>
</p>
</td></table>
</ol>
</li>
<h2><li>Microscope Set-up and Kohler Illumination:</h2>
<ol>
<li>This experiment will be performed using bright-field microscopy. Ensure the ring on the
condenser diaphragm is set at 0 (not PH 1 or PH 2), because the phase ring at the PH1 or PH2
setting attenuates the light.</li>
<li>Once your specimen slide is prepared, bring it to the microscope and focus on the specimen
under the x10 objective.</li>
<li>Next close the field diaphragm all the way shut so you can see the edges, they may appear
blurry.</li>
<li>Use the condenser focus knob to bring the edges of the field diaphragm into the best possible
focus.</li>
<li>Next use the condenser-centering screws to bring the closed field diaphragm into the center of
the field of view.</li>
<li>Then open the field diaphragm such that it is slightly larger than the field of view.</li>
<li>You are now set up for Kohler illumination.</li>
</ol>
</li>
<h2><li>Light Treatments</h2>
<p>The wavelength dependence of light on chloroplast translocation will be investigated. From the
absorption spectrum of chlorophyll shown in Figure 1, it is known the chlorophyll strongly absorbs both
blue and red light, thus for the wavelength experiments we will be using blue and red filters band-pass
filters, their transmissions are shown in Figure 4. The blue band-pass filter emits light at a peak
wavelength of 441nm ± 10nm. The red filter is a band-pass filter with a tail at longer wavelengths. This
filter has a peak wavelength of 623nm ± 45nm.</p>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig4.png|450px|left]]
<b>Figure 4: Emission Spectra of Band-pass Filters:</b>The blue bandpass
filter was fit to a
Gaussian function and the
red band-pass filter was fit
to a Lorentzian Function to
account for the long tail
end. Both fit functions are
shown in black.
<br clear=right>
</p>
</td></table>

<p>The cells will be illuminated with both blue and red light to determine the wavelength dependence of
light on chloroplast translocation as described in the following outline:</p>
<ol>
<li>Take the blue band-pass filter labelled 441nm and place it above the light source under the
condenser diaphragm.</li>
<li>Double click on the Desktop folder called <b>BPHS_4090_Viridis_Systrophe</b>, and then open the
Excel spreadsheet called <b>“Photon_Flux_Calculator.xls”</b>. This spreadsheet is already
programmed to do all the difficult calculations for you to save you time in the lab!</li>
<li>With the blue filter in place turn on the radiometer, use the probe to measure the output power
from the light source; this is done by removing the sample slide and placing the probe in the
light path where the objective is. You may have to remove one of the eyepieces to get the
probe in the light path, do this carefully and ensure you leave the x10 objective (labelled Fluor
10) in place, since this will be used for the light treatments. Ensure the probe is collecting all the
light. You should measure a power reading in microwatts; enter this into the spreadsheet to get
a photon flux as an output in units of μmol m<sup>-2</sup> s<sup>-1</sup> as shown in Figure 5.</li>

<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig5.png|650px|left]]
<b>Figure 5:</b> The flux calculator has separate sections for the red and blue bandpass
filters. The cells will feel different energies corresponding to the different wavelengths of light
according to the Planck relationship <i>E = hc/λ </i>.
<br clear=right>
</p>
</td></table>

<li>For the wavelength experiment a photon flux of around <b> 200μmol m<sup>-2</sup> s<sup>-1</sup></b> is ideal to observe the
effect of chloroplast translocation. Adjust the voltage dial knob on the microscope until you get
a power reading which gives you a photon flux close to this value,<b>once you do so, record your
input power.</b></li>
<li>Next place the cell sample back in the field of view, drop the neutral density filter labelled ND 2
in the light path to avoid pre-treating the cells with light. You’ll have to adjust the microscope
again for Kohler Illumination, but it shouldn’t take much adjusting at this point.</li>
<li>Move the microscope stage to capture as large of a cell population as possible, try to aim for 20
or more cells if possible by looking through the eyepiece as you move the stage.</li>
<li>Ensure your cell sample is in good focus, you’re now ready to make a movie of chloroplast
translocation.</li>
</ol>

<h2><li>Making a Movie of Chloroplast Translocation:</h2>
<ol>
<li>Turn on the CoolSnap CCD camera.</li>
<li>Open MicroManager 1.3 by double clicking the icon on the desktop, you’ll notice the program
ImageJ (Figure 6) opens as well, this will be used later for image processing.
Figure 6 – MicroManager Software</li>
<table width=600 align=center><td>
<p align=justify>[[File:Ct_fig6.png|550px|left]]
<b>Figure 6:</b> MicroManager Software.
<br clear=right>
</p>
</td></table>
<li>Click on Live Feed and turn the eyepiece mounting block about 45 degrees to the left, this allows
light to pass through to the CCD camera and will project a live image onto the computer
monitor.</li>
<li>Next lift up the ND2 filter and adjust the exposure time, start by setting it to 1ms, you’ll also
notice the cells on the screen are out of focus, use the fine focus adjustment until you get crisp
edges along the periphery of the cell.</li>
<li>Adjust the exposure until you get a live image that isn’t oversaturated (Figure 7).</li>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig7.png|650px|left]]
<b>Figure 7:</b> MicroManager Live Feed.
<br clear=right>
</p>
</td></table>
<li>Due to the limited field of view, you’ll only be able to get about 1 cell in the live feed. Move the
microscope stage slowly (since the response from the CCD is slightly delayed) until you get 1 cell
in the field of view as shown in Figure 7.</li>
<li>Click on “Multi-D acq.” Then set the number to 360 and interval to 10 seconds, and then hit
acquire (Figure 8). This tells the program to take an image every 10 seconds; this will eventually
give you a movie 60 minutes long, which should allow you to see some significant chloroplast
translocation as shown in Figure 2.</li>
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig8.png|450px|center]]
<b>Figure 8:</b> Multi-Dimensional Acquisition.
<br clear=right>
</p>
</td></table>
<li>Once finished click on the save icon. Save your data in the directory
<b>C:\BPHS4090_Viridis_Systrophe, label your data by your name, date and title of work.</b> This
will create a file folder with all the images you acquired.</li>
<table width=300 align=center><td>
<p align=justify>[[File:Ct_fig9.png|250px|center]]
<b>Figure 9:</b> Sequence Options.
<br clear=right>
</p>
</td></table>
<li>Next you’ll be using ImageJ. Click “File->Import->Image
Sequence”, double click on your file folder and select
one of the images and press Open. Another window
will open called “Sequence Options” (Figure 9), it will
verify the number of images you have and open them
as a stack, make sure the box titled “Sort Names
Numerically” is checked and click Ok.</li>
<li>This may take a little while, once all the images are
loaded, use the mouse cursor icon to make a box
around your cell and click on “Image->Crop”. Every
image you imported will be cropped.</li>
<li>Next click on “Image->Adjust->Brightness/Contrast”,
another window called “B&C” will open; adjust the
settings to ensure your images look the best they can.
Once happy with the results click on Set, a new window
will open, make sure the “Propagate to all open
images” box is checked and click OK (Figure 10). This
will adjust the brightness and contrast for every image
in your imported stack.</li>

<table width=400 align=center><td>
<p align=justify>[[File:Ct_fig10.png|350px|center]]
<b>Figure 10:</b> Adjusting Image Brightness and Contrast.
<br clear=right>
</p>
</td></table>
<li>Once your image is adjusted for proper brightness and contrast select “File->Save as->AVI”,
select a frame rate of about 10fps, this will make a movie 36 seconds long assuming you took
360 frames. A movie at this speed will dramatically show chloroplast translocation assuming the
one cell you chose to view undergoes the effect. Save the AVI in the directory
<b>C:\BPHS4090_Viridis_Systrophe by your name, date and title of work.</b></li>
<li><b>Copy your movie and the file called “Viridis_Systrophe_Spectrum.xls” onto a USB Key, this
spreadsheet will be used for one of the questions. You can find the file in the desktop folder
called “BPHS_4090_Viridis_Systrophe”.</b></li>
<li>Look at your cell sample through the field of view and record how many cells are incipient
systrophe and how many are complete systrophe using the criteria in Figure 2.</li>
<li>Once this is done discard your cell sample slide in the bin labelled “Sharps”</li>
<li>Next prepare a new sample slide and repeat the above procedure using the red filter labelled
623nm. DO NOT make a movie using the red filter; record the total number of cells in the field
of view and illuminate the cells for an hour. After the light treatment record the number of
incipient and complete systrophe. Be sure to adjust your power to get the desired photon flux
of 200 μmol m<sup>-2</sup> s<sup>-1</sup> and ensure your microscope is set up for Kohler illumination.</li>
</ol>
</ol>

<h1> Questions/Write-up </h1>

<p>Answer the following questions in your lab notebook.</p>
<ol>
<li>An additional experiment was performed [8] by illuminating only half the cell at a time. The
results of this experiment are shown below:
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig11.png|700px|center]]
<b>Figure 11:</b> Half Cell Illuminations: This particular cell was illuminated under broad band-pass
irradiation with a photon flux of <b>1500μmol m<sup>-2</sup> s<sup>-1</sup>.</b> After an hour of irradiation, chloroplasts have
vacated the illuminated area.
<br clear=right>
</p>
</td></table>
<p>What can you conclude about these results? Is chloroplast translocation a protective
mechanism for the cell or rather an avoidance mechanism? Explain in a few sentences.</p></li>

<li>Imagine you had a source of UV light available and you wished to perform an additional
experiment by illuminating the cells with UV light as you did earlier with the blue and red bandpass
filters. Assume your UV light source emits at a peak wavelength of 250nm ± 10nm. Taking
into account the absorption spectra of chlorophyll shown in Figure 1 and that DNA absorbs light
at 260nm what would you expect to happen?</li>

<li>During this lab we examined the wavelength dependence of light on systrophe, however other
experiments [8] demonstrated there is also an intensity dependence as shown in the Figure
below:</li>

<table width=500 align=center><td>
<p align=center>[[File:Ct_fig13.png|500px|center]]
<b>Figure 13:</b> Intensity Dependence of Light on Incipient and Complete Systrophe.
<br clear=right>
</p>
</td></table>

<p>At a photon flux of around <b>2000μmol m<sup>-2</sup> s<sup>-1</sup></b> there is a dramatic increase in the number of
incipient systrophe. Remember incipient means the cells are in the process of undergoing
chloroplast translocation. The photon flux we receive from the sun is around <b>2000μmol m<sup>-2</sup> s<sup>-1</sup></b>.
What does this imply?</p></li>

<li>Photosynthetic organisms use wavelengths between 400nm and 800nm for photosynthesis.
From Figure 1, it can be seen that chlorophyll strongly absorbs blue and red light. Taking this
into account and your answers to questions 1 and 2 what role do you think systrophe plays in
photosynthesis? Are they related? Please discuss in a few sentences providing insights into
your thinking process.</li>
</ol>

<p>For the formal write-up for this experiment, briefly describe the purpose of the experiment, the
experimental set-up and the physics behind the biological process we’re observing, this will be your
introduction. You should type up your results into a table comparing the illumination trials using red
and blue filters. Provide a brief discussion interpreting your results. In addition you need to provide
answers to the following questions.</p>
<ol>
<li>The effect of systrophe is thought to be a defence mechanism for the cell by decreasing the
absorptive cross sectional area of chloroplasts within the cell. We can test this hypothesis by
using a simple mathematical model to represent the cell. Assume each cell of E. viridis in its
resting state is a sphere with a shell of chloroplasts along the periphery of the cell. In reality
chloroplasts are also distributed radially along the cytoplasmic strands within the cell (analogous
to the spokes of a bicycle wheel); however for simplicity we will only consider the outer shell of
chloroplasts. After the cell undergoes systrophe, the chloroplasts surround the nucleus in a
thicker shell. In the table below some data are provided of cellular dimensions for 3 arbitrary
cells before and after systrophe using time lapsed images <ref>Web-published: http://www.yorku.ca/planters/student_reports/viridis_systrophe.pdf (accessed
9 September 2010).</ref>.

<table width=500 align=center>
<tr><td><p align=center><b>Table 1:</b> Cellular Dimensions before and after Systrophe.</p></td></tr>

<tr><td><table width=500 align=center border>
<tr><td></td><td>Cell 1</td><td>Cell 2</td><td>Cell 3</td></tr>
<tr><td>Cell Diameter (μm)</td><td>122.5</td><td>108.5</td><td>130.5</td></tr>
<tr><td>Systrophe Diameter (μm)</td><td>80.1</td><td>64.7</td><td>103.8</td></tr>
<tr><td>Nucleus Diameter (μm)</td><td>41.8</td><td>45.8</td><td>61.3</td></tr>
<tr><td>Thickness of Chloroplast Shell
(Before Systrophe) (μm)</td><td>4.6</td><td>5.5</td><td>5.3</td></tr>
</table>

</td></tr></table>

<p>Use the data in Table 1 as well as the physical properties of the cells included in the
introductions and the Beer-Lambert Law (Equation 1.3) to fill in the table below:</p>
<table width=150 align=center><td>
<p align=justify>[[File:Ct_eqn1_3.png|150px|center]]
<br clear=right>
</p>
</td></table>
<p>Where I is the transmitted intensity of light passing through the cell, Io is the initial intensity your
light source, ε is the extinction coefficient of chlorophyll, C is the concentration of chlorophyll
and l is path length which the light passes through.</p>
<table width=700 align=center><td>
<p align=center><b>Table 2: </b> Absorptive Properties of Cells before and after Systrophe.
[[File:Ct_tab2.png|650px|center]]
<br clear=right>
</p>
</td></table>

<p>Based on your results in Table 2, does systrophe provide a protective mechanism for the cell by
decreasing the absorptive cross sectional area? Why would an increase in absorption
potentially be damaging for the cell? Consider both absorption and cross sectional area.</p>
</li>

<li>Open the spreadsheet titled <b>“Viridis_Systrophe_Spectrum.xls”</b>; this is an emission spectra of
the tungsten halogen bulb used for illuminating the cell. Calculate the total energy the cells
would receive from the whole spectrum of light emitted from this source by normalization. You
can use the spreadsheet to do your calculations and may wish to attach it to your lab report.</li>
<li>Below is the emission spectrum from the tungsten halogen bulb used for illuminations. From
the Planck Blackbody Distribution Law derive Wien’s Displacement Law which relates the colour
temperature of a blackbody to the peak wavelength it emits at. If we treat tungsten as a
blackbody, what is the color temperature associated with the peak wavelength observed in
Figure 12? Does this make sense considering the melting point of tungsten is about 3350K?
What correction do we need to take into account when deriving Wien’s Displacement Law from
Planck’s Blackbody Distribution Law to account for the unusual high melting point implied from
Figure 12?</li>

<table width=500 align=center><td>
<p align=center>[[File:Ct_fig12.png|500px|center]]
<b>Figure 12:</b> Tungsten Halogen Emission Spectrum.
<br clear=right>
</p>
</td></table>

<li>During systrophe chloroplasts are translated across the cell by molecular motors. It isn’t known
exactly which molecular motors are responsible for chloroplast translocation, however it should
be noted chloroplast translocation can be inhibited when cells of <i>E. viridis</i> are treated with 2,4 –
Dinitrophenol (DNP) [4]. The structure of DNP is shown in Figure 14.

<table width=400 align=center><td>
<p align=center>[[File:Ct_fig14.png|200px|center]]
<b>Figure 14:</b> Structure of 2,4 Dinitrophenol.
<br clear=right>
</p>
</td></table>
DNP acts as an uncoupling agent by disrupting the proton gradient the cell uses to create ATP.
Any energy the cell would have gained from ATP production is lost as heat. ATP provides the
energy for molecular motors to shuttle organelles across the cell. Based on the structure of
DNP, taking into account hydrophilicity and hydrophobicity and any partial charges on the
molecule explain how DNP inhibits chloroplast translocation. Does this provide any insight into
the energy mechanisms involved during systrophe?
</li>

<h1>References</h1>
<References/>

File:Fluorescence scans.png

2012-08-10T11:30:26Z

Rlew: Fluorescence scans obtained with a fluorometer in the teaching labs for Photosynthesis (SC/BIOL 4160) [http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/BIOL_4160]

Fluorescence scans obtained with a fluorometer in the teaching labs for Photosynthesis (SC/BIOL 4160) [http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/BIOL_4160]

Main Page/BPHS 4090/Optical Tweezers of Onions

2012-07-28T16:57:12Z

Rlew:

<h1> Introduction </h1>

<h2> The Physics of Optical Tweezers </h2>
<p>The principle of optic tweezers was first proposed and discovered by Arthur Ashkin in 1970.<ref> Ashkin, A. (1970). "Acceleration and Trapping of Particles by Radiation Pressure". Phys. Rev. Lett. 24: 156–9. doi:10.1103/PhysRevLett.24.156. http://prola.aps.org/abstract/PRL/v24/i4/p156_1.</ref> The principle relies on the refraction of light as it passes between media of differing indicies of refractions (symbolized by <i>n</i>), and the Gaussian profile of a trapping laser beam which is most intense at the central axis. Recall that a photon with wavelength λ and moving along the z-axis has of momentum equal to:
<table width=80 align=center><td>
<p align=justify>[[File:ot_eqn1.png|80px|center]]
<br clear=right>
</p>
</td></table>
Note this this value is very small, and for large objects, the force from reflecting a photon is very small. However, for a large flux of photons (such as a from a laser) spread over a small area (focuss by powerful microscope objective) one can exert a meaning force on a small particle (such as micron-sized beads, or intercellular particles) As a photon passes through a transparent sphere of greater index of refraction than the surrounding medium, it will refract (change direction). This results in a change in momentum. As a reaction to this change in momentum, momentum is imparted to the bead in the opposite direction. The ray diagram for this effect is given in Figure 1. It is important to realize that the magnitude of the refraction of the photon depends on the difference of the indicies of refraction of the transparent sphere and surrounding medium. </p>

<table width=400 align=center><td>
<p align=justify>[[File:Ray_diagram.png|400px|border|center]]
<b>Figure 1 -</b> In part a), there is more laser intensity on the right of the bead due to the gaussian profile of the laser beam, and hence there is a net force to the right. In part b), the bead is centered and the net force in the left/right direction zero.
<br clear=right>
</p>
</td></table>

<p>If the particle to be trapped is not so spherical, or scatters photons from the laser beam rather than refracts the photons, there will be a strong force pushing the sphere along the axis of laser propogation- like a water hose pushing a beach ball. It is imperative that the object to be trapped allows most of the light to pass through it, rather than bounce off its surface. You may have seen an example of optical trapping of latex beads in PHYS4061.</p>

<h2> Structure of an Onion Cell </h2>
<p>Plant cells have the general properties of a rigid cell wall, a large open vacuole, a nucleus, and cytoplasm containing organelles in the spaces between the cell walls and vacuole. The onion cell is a classic and often-used example of this structure. The organelles in the cytoplasm are small (between 0.5 and 1 micron), and roughly spherical in nature- prime candidates for optical tweezing with a laser. Organelles move throughout the cyctoplasm either along action filaments, and along the the endoplasmic reticulum network.</p>

<table width=400 align=center><td>
<p align=justify>[[File:400px-Onioncell.gif|400px|border|center]]<ref> Source: N. S. Allen and D. T. Brown, 1988. Dynamics of the Endoplasmic Reticulum in living onion epidermal cells in relation to microtubules, microfilaments, and intracellular particle movement. Cell Motility and the Cytoskeleton 10:153-163</ref>
<b>Figure 2: The onion cell</b>
<br clear=right>
</p>
</td></table>

<p> The organelles move throughout the onion cell only in the cytoplasm and not through the vacuole. There are strands of cytoplasm located sporadically thoughout the cell. These strands themselves can be seen by the imagining optics when focused appropriately.
Within these strands of cytoplasm are actin fibers along which organelles are transported. The process by which this occurs is demonstrated nicely in the following animation.
[http://multimedia.mcb.harvard.edu/anim_myosin.html]. Movies of the streaming can be viewed at the following link [http://www.yorku.ca/planters/movies.html#Onion]</p>

<h2>Preparation of onion epidermis for optical trapping experiments <ref>Diagrams from Peterson LR, CA Peterson, and LH Melville (2008) Teaching Plant Anatomy through Creative Laboratory Exercises. National Research Council of Canada. Page 17.</ref> </h2>
<p>The common onion (<i>Allium cepa</i>) is often used to examine individual plant cells because of the ease of isolating sheets of cells that are one cell thick. The onion bulb can be sectioned into quarters or eighths. Then, the individual scale leaves can be separated.</p>

[[ File:Onionprep_1.jpg|270px]]
[[ File:Onionprep_2.jpg|270px]]
[[ File:Onionprep_3.jpg|270px]]

<p>The exposed concave surface can be scored with a sharp razor blade. With a very fine pair of forceps, pieces of the epidermis can be lifted (note the transparency of the peel). Before doing so, have a microscope slide ready with a drop of distilled water (or artificial pond water) so that the peel doesn’t become dehydrated. Unlike the photographs, a thin strip of Vaseline will be placed on the microscope slide in a rectangular shape slightly smaller in dimensions then the cover slip. Having placed the epidermal peel in the water inside the Vaseline ‘dike’, carefully (gently) place the coverslip on top, pressing to create a seal around the perimeter.</p>
[[ File:Onionprep_4.jpg|270px]]
[[ File:onion3.jpg|270px]]
[[ File:Onionprep_6.jpg|270px]]
<p>Now ready to place in the holder on the optical bench (left), here is what the cells will look like (right, the nucleus and transvacuolar strands are indicated).
[[ File:onion8.jpg]]
</p>

<h1> Lab Exercise: Optical tweezing of onion cells </h1>

<h2> Apparatus </h2>
<p> Although the apparatus used in this experiment may look unfamiliar and unlike anything you have used, closer inspection reveals that the system is just an open microscope mounted on its side. All basic components of microscope are laid out on the optics table- the objective, the sample manipulator, the light source and condenser, interacting light source, and the CCD camera. Since you are very familiar with details of a microscopy, this apparatus shoudln't prove intimidating. Instead of you interacting light source being used for fluorescence, you will use a powerful, monochromatic light source to interact with the sample.</p>
<p> Of course, this powerful light source is a laser. Solid-state lasers have made high-power monochromatic light cheap and available. In the recent past, one would have required a 1.5-meter long laser tube, with a power supply the size of an small filing cabinet to have the equivalent performance as the thumb-sized laser we will use. Figure 3 shows a general overview of the apparatus, with the major components indicated.</p>

<table width=600 align=center><td>
<p align=justify>[[File:Ot_fig3_overview.jpg|600px|border|center]]
<b>Figure 3 -</b> Overview of the Apparatus
<br clear=right>
</p>
</td></table>

<p> The laser/optical system is set up to ensure that the focus of the laser light through the objective is co-planar with the focus of the imaging optics of the CCD. The laser power is controlled by the combination of a1/2-λ plate and polarizing cube (see Figure 4). Light generated by the laser is mostly polarized along one axis. The 1/2-λ plate rotates this axis of polarization. The polarization cube separates light out into vertical and horizontal polarized light. Hence, at one position of the half-wave plate, the light will be predominately at the polarization required to pass straight through the cube. At 90 degrees to that rotation of the 1/2-λ plate, the light will be predominantly of the polarization to be directed out the side of the cube. Therefore, rotating the 1/2-λ plate will serve as our method of varying the laser power. In order to measure the laser power, a beam sampler is placed after the polarizing cube, and directs a fixed percentage of the laser beam to the side for measuring on a photodiode detector.</p>
<p> Once the laser power is controlled and sampled, it passes through a lens which adjusts the divergence of the laser beam such that the co-planar focus condition mentioned above is set. Following this, the laser is relfected into the 100 DIN oil objective off of a dichroic mirror. The dichroic mirror has the property of being a very strong relfector of 532nm, but a very weak reflector at shorter wavelengths. Therefore, light from the lamp used for imaging can pass the opposite way through the dichroic mirror and produce an image on the CCD camera. </p>

<table width=600 align=center><td>
<p align=justify>[[File:Ot_fig4_optical_path.jpg|600px|border|center]]
<b>Figure 4 -</b> The optical system.
<br clear=right>
</p>
</td></table>

<p> The laser is a DPSS (Diode-Pump-Solid-State) laser at 532nm rated to provide 100mW at 3V. Due to its power, it is classified as a Class IIIB laser. The procedure for operating the laser Figure 5) is as follows:</p>
<table width=400 align=left><td>
<p align=justify>[[File:Ot_fig5_laser.jpg|370px|border|left]]
<b>Figure 5 -</b> Laser control system.
<br clear=right>
</p>
</td></table>
<ol>
<li> Put on the safely goggles. </li>
<li> Turn on the mulitmeter to measure DC current, on the 10A scale. </li>
<li> Turn the key 90 degrees clockwise on the laer power supply.</li>
<li> Enure the middle knob of the power supply is fully counterclockwise.</li>
<li> Please the toggle switch in the "down" position.</li>
<li> Slowly turn the middle knob clockwise while monitoring the current on the meter. You should be able to visibly see the laser on when the current increase above 0.18A. UNDER NO CIRCUMSTANCES ALLOW THE CURRENT TO INCREASE PAST 0.45A.</li>
</ol>

<br clear=all>

<p> The microscope slide is mounted on a 3-axis precision translation stage (See Figure 6). The two axes transverse to the laser have piezo-controlled actuators which are controlled from the computer. All three axes can also be controlled by the coarse- and fine-adjust knobs located on the translation stage.</p>

<table width=400 align=center><td>
<p align=justify>[[File:Ot_fig6_stage.jpg|400px|border|center]]
<b>Figure 6 -</b> The Translation Stage.
<br clear=right>
</p>
</td></table>

<h2> Computer Control </h2>

<p> A LabView program is used to view the output from the CCD camera, and also to control the X-Y position of the sample. The layout of the user interface is described below.</p>
<table width=700 align=center><td>
<p align=justify>[[File:Optical_tweezers_vi.png|700px|border|center]]
<br clear=right>
</p>
</td></table>

<h2> Calibrate Optical Trap Depth </h2>
<p> The goal of this experiment will be to determine the stength of the myosin motors which move the spherosomes along actin filaments. In order to achieve this goal you must first calibrate how strongly the spherosomes are trapped in the optical tweezers as a function of laser power. For this, we will use spherosomes moving in cytoplasmic medium near the cell walls, but not along actin fibers. The spherosomes are moving through the cytoplasm, adjacent to the endoreticulum network. Cell cytoplasm is a very complicated component of the cell, and can be characterized as a non-Newtonian fluid. The viscosity of the cytoplasm depends on the shear applied to the cytoplasm. For our purposes, we will take an educated guess of a viscosity of 20 times that of the viscosity of water. To determine the trapping force on the spherosome, we will move the spherosome through the cytoplasm as a constant velocity and see which velocity is great enough to dislodge the spherosome from the trap.
The force exerted by the viscous cyctoplasm on a sphere is<ref>S. Henon, G. Lenormand, A. Richert, F. Gallet, Biophysical Journal <b>76</b> 1145 (1999)</ref>:

<table width=150 align=center><td>
<p align=justify>[[File:ot_eqn2.png|130px|left]]
<br clear=right>
</p>
</td></table>

Where <i>R</i> is the radius of the sphere, <i>η</i> is the viscosity of the cytoplasm, and <i>v</i> is the velocity of the sphere through the cytoplasm. </p>

<p>
In order to have nice repeatable motion with the piezo translators, it is necessary to start at 0V applied to the piezo controlling the direction in which you want to move.
</p>

<h3> Method</h3>
<ol>
<li>Determine the position of the laser. Notice that you can as you move the onion slide towards the microscope objecive using the z-xis manual control, the cover slip will come into focus first, and the laser position will be visible due to scattered light off the coverslip. Using the "region of interest" marker tool, place a ~2 micron circle around this position. </li>
<li>Focus on the top cell wall. As you continue moving the onion slide closer to the microscope objective, first the top cell wall will come into focus, then the middle parts of the cells, then, if the sample isn't too thick, the lower cell wall. You will see lots of organelles moving in many directions.</li>
<li>Adjust the half-wave plate for maximum transmission.</li>
<li>Trap one of these spherosomes in the optical trap.</li>
<li>Move at constant velocity using the velocity control panel. Note that the actual velocity is also displayed and you should use this in your calculations. If you are having a problem having the actual velocity match with the set velocity, reduce the number of steps. Conversely, if the velocity movement seems jumpy, increase in the number of steps. </li>
<li>Adjust the laser power by rotating the half-wave plate. Repeat above step, and make a table of relative laser power and minimum velocity required to dislodge the spherosome from the trap.</li>
<li>From the above table calculate the force on the spherosome, and plot the data on a graph.</li>
</ol>

<p>You now have a calibration graph that you can use in the next section to read off the force exerted on a spherosome for a given setting of the half-wave plate.</p>

<h2> Determine Strength of Myosin Motor </h2>

<p> Now, rather than trapping spherosomes moving near the cell wall, you will see what force is required to trap spherosomes moving along actin fibers.</p>

<ol>
<li> Identify an actin fiber located in the middle of the cell. They will appear a dark tracks moving through the middle of the empty vacuole. Ensure the actin fiber is active and has organelles moving along it.</li>
<li> Adjust the laser power to minimum using the half-wave plate, and place the laser focus on a part of the actin fiber. Ensure that system is focused such that spherosomes come into focus as the pass through the laser trap.</li>
<li> Are spherosomes affected at all by the laser beam? Do they scatter photons? Are they stopped?</li>
<li> Adjust the half-wave plate so more laser power is available for trapping. And answer above questions. Record data for you observations in a chart.</li>
<li> Repeat above steps until you reach full laser power</li>
<li> Repeat above steps for other actin fibers.</li>
</ol>

<h1> Questions and Discussion </h1>

<ol>
<li> A 532nm green laser with a power of 100mW passes from air into a flat glass block placed at Brewster's angle. The laser is polarized such that all photons are refracted, and none are reflected. What is the force exerted on the glass in the transverse direction? </li>
<li> How would the strength of the optical trap change if purple light were used rather than green light? Wat are some possible concerns with changing the wavelength?</li>
<li> Describe how a 1/2-λ plate works.</li>
<li> Would you expect the optical trap depth for a spherosome on the top cell wall would change to a spherosome in the middle of the cell? Why?</li>
<li> Comment on your results from attempts to determine the strength of the myosin motor force. </li>
</ol>

<h2>References</h2>

<references/>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-07-13T16:35:36Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://eol.org/pages/11583/overview)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie. The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-07-13T16:33:46Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://www.eol.org/pages/11583)(black)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie. The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-06-19T17:38:17Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://www.eol.org/pages/11583)(black)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)(red)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)(blue)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie. The movie is available at http://www.yorku.ca/planters/movies.html#Chara. Detailed notes on culturing Chara are provided at the following <b>BioWiki</b>: http://biologywiki.apps01.yorku.ca/index.php?title=Main_Page/Culturing_Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-06-14T17:23:24Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://www.eol.org/pages/11583)(black)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)(red)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)(blue)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=1000 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|1000px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie. The movie is available at http://www.yorku.ca/planters/movies.html#Chara.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

File:Chara Growth montage.png

2012-06-14T17:20:00Z

Rlew:

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2012-06-14T17:19:42Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://www.eol.org/pages/11583)(black)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)(red)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)(blue)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>
<table width=700 border=1 align=center><td>
<p align=justify>[[File:Chara_Growth_montage.png|700px|right]]
<b>Growth of Chara</b> The montage is snapshots from a time lapse movie.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2011-12-15T17:46:11Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971)<ref>McCombie, A.M. and I. Wile (1971) Ecology of aquatic vascular plants in southern Ontario impoundments. Weed Science 19:225–228.</ref>. It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://www.eol.org/pages/11583)(black)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)(red)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)(blue)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2011-12-15T17:42:09Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971). It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954)<ref>Collander, R. (1954) The permeability of Nitella cells to non-electrolytes. Physiologia Plantarum 7:420–445.</ref>. He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://www.eol.org/pages/11583)(black)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)(red)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)(blue)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

Talk:Main Page/BPHS 4090/microscopy II

2011-11-29T21:23:06Z

Rlew: Protected "Talk:Main Page/BPHS 4090/microscopy II" ([edit=autoconfirmed] (indefinite) [move=autoconfirmed] (indefinite))

Talk:Main Page/BPHS 4090/microscopy II

2011-11-29T21:22:56Z

Rlew: Blanked the page

Talk:Main Page/BPHS 4090

2011-11-29T21:22:26Z

Rlew: Protected "Talk:Main Page/BPHS 4090" ([edit=autoconfirmed] (indefinite) [move=autoconfirmed] (indefinite))

Talk:Main Page/BPHS 4090

2011-11-29T21:22:19Z

Rlew: Blanked the page

Talk:Main Page

2011-11-29T21:21:55Z

Rlew: Protected "Talk:Main Page" ([edit=autoconfirmed] (indefinite) [move=autoconfirmed] (indefinite))

Talk:Main Page

2011-11-29T21:21:43Z

Rlew: Blanked the page

Talk:Main Page

2011-11-29T14:31:45Z

Rlew: Blanked the page

Talk:Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2011-11-28T22:38:05Z

Rlew: Protected "Talk:Main Page/BPHS 4090/ElectroPhysiology of Chara revised" ([edit=autoconfirmed] (indefinite) [move=autoconfirmed] (indefinite))

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2011-11-25T18:45:10Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:impalement_examples.png|300px|right]]
<b>Figure 1.6: Examples of Cell Impalements.</b>
Photos of various cells are shown on the left (A is an algal cell, B is a fungi, C is a plant cell). Examples of the electrical changes that occur upon impalement of the cell (and upon removal of the microelectrode) are shown on the right. Electrical traces for impalement into <i>Chara</i> will be similar.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971). It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954). He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://www.eol.org/pages/11583)(black)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)(red)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)(blue)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>

File:Impalement examples.png

2011-11-25T18:41:57Z

Rlew:

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2011-11-25T18:30:40Z

Rlew:

<h1>Overview</h1>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_1.png|500px|right]]
<b>Workstation for Electrophysiology.</b> The various components of the electrophysiology workstation are shown here and are described in the lab exercise. The Faraday cage shields the measuring apparatus from external electromagnetic noise. The micromanipulators, with xyz movement, are used to align the micropipette (mounted on the headstage), then descend to the cell to effect impalement, all viewed through the dissecting microscope. Note that the entire apparatus is mounted on an optical bench, to isolate the system from mechanical vibration.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Overview_2.png|300px|right]]
<b>Specimen Chamber, Micropipette Holder and Micropipettes.</b> The internodal cell is centered in the specimen chamber (filled with APW7) and gently held down by weights on either side of the observation window. Micropipettes should be filled with 3 M KCl as described in the lab exercise, inserted into the pre-filled (with 3 M KCl) micropipette holder and secured by tightening the knurled screw. See the lab exercise for details.
<br clear=right>
</p></table>

<table width=800 border=1 align=center><td>
<p align=justify>
<b>Example of Single and Dual Microelectrode Impalements into <i>Chara australis</i>.</b> The cell was first impaled with one microelectrode, then impaled with two. Note that the voltage after two impalements eventually reached about -200 mV (not shown). The action potential has not been confirmed, but exhibits the characteristic duration and shape of an action potential in Chara.[[File:01_Overview_3.png|780px|center]]
<br clear=right>
</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew with assistance from Dr. Matthew George and Israel Spivak. 26 may 2011.</ref>: The Electrical Properties of ''Chara''<ref>I would like to acknowledge the advice and assistance of Professor Mary Bisson (University at Buffalo), who has researched the electrophysiology of ''Chara corallina'' for many years. Her advice and generous donation of ''Chara'' cultures made this lab exercise for York Biophysics students possible.</ref> </h1>

<p>
In this exercise, you will have the opportunity to measure the electrical properties of a single cell. The organism you will be using is a giant ''coenocytic'' (multi-nucleate) internodal cell of the green alga ''Chara australis''. This is a common model cell for electrical measurements, and has been used extensively by biophysicists for more than 50 years. Its large size makes the challenges of intracellular recording simpler, allowing the researcher to focus on more sophisticated aspects of cell electrophysiology.</p>

<h2> Objective </h2>
<p>To measure the electrical properties of the plasma membrane of a cell: 1) Voltage measurement (single microelectrode impalement); and 2) Action potentials.
</p>
{||123||345||456||}

<h2> Introduction </h2>
<p>You are already aware of the electrical properties you will be measuring, at least in the context of electronics. In electrophysiology, electron flow is replaced by ion flow. The basic electrical parameters of the cell are shown in Figure 1.1.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.1.png|400px|right]]
<b>Figure 1.1: Electrical circuit of an internodal cell of Chara.</b> The diagram shows the electrical network of the plasma membrane (a capacitance, C, in parallel with a voltage, V, and resistance R.
Note that both the cytoplasm inside the cell and the external medium have an electrical resistance. This can complicate electrical measurements, but in your exercise, the resistance of the membrane is
much greater than cytoplasm and external medium resistances so their effect can be ignored. The electrical properties (V, I, C and R) are related through the basic equations: V = IR and I = C(dV/dt).
In this exercise, you will be measuring voltage, resistance and capacitance.
<br clear=right>
</p>
</td></table>

<p>There are multiple resistive (and capacitative) elements in the circuit. A brief reminder of how resistances and capacitance in series and in parallel behave is shown in Figure 1.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.2.png‎|200px|right]]
<b>Figure 1.2: Resistances and Capacitances</b> The diagrams should remind you of the behaviour of resistance and capacitance when they are connected in series or in parallel. Both situations arise in the case of measurements of the electrical properties of biological cells (Fig. 1.1). Experimentally, resistance (R<sub>total</sub>) can be measured by injecting a known current into the cell, the steady state voltage change is related to R<sub>total</sub> by the equation V=IR. For biological cells, the standard units are (mV) = (nA)(MΩ). Capacitance is determined by measuring the voltage change over time while injecting a current pulse train into the cell. The shape of the voltage change is exponential, V<sub>t</sub>=V<sub>t=0</sub>• exp(–t/(RC)), eventually reaching steady state. If R is known, then C can be determined. The plasma membrane resistance and capacitance need to be normalized to the area of the cell membrane (area specific resistance: Ω cm<sup>2</sup>; specific capacitance per area: F cm<sup>–2</sup>). Note that the units match the behavior of resistances and capacitance in parallel. That is, a larger cell has greater membrane area. There is more ‘resistance in parallel’, and the total resistance (Ω) is lower. Conversely, a larger cell has a greater capacitance (F).
<br clear=right>
</p>

</td></table>

<p>Of course, like all good things, resistive network analysis can be taken too far (Fig. 1.3).</p>

{|border="1" width="700" align="center"
|<b>Figure 1.3: Resistive network analysis can be taken <i>way</i> too far.</b> As shown in this cartoon from Randall Munroe (http://xkcd.com/356/) entitled “Nerd Sniping”.
[[File:01_Figure_1.3.png|600px|center]]
|}

<h2> Methods </h2>
<p><b>Microelectrode preparation and use.</b> You will be provided with pre-pulled micropipettes. The micropipettes are made from borosilicate glass tubing, and pulled at one end to form a very fine and sharp tip. The dimensions at the tip are an outer diameter of about 1 micron (10<sup>-6</sup> m), an internal diameter of about 0.5 micron. The micropipette contains a fine glass filament, bonded to the inner wall of the glass tube. This provides a conduit (via capillary action) for movement of the electrolyte filling solution right to the tip of the micropipette. A more detailed description of micropipette fabrication and physical properties is presented in Appendix I.</p>
<p>To fill the micropipette, insert the fine gauge needle at the back of the micropipette so that the needle tip is about 1 cm deep (Figure 1.4). Fill the back 1 cm of the glass tube with the solution (3 M KCl) and carefully place the micropipette on the support provided. Due to capillary action, the micropipette will fill all the way to the tip (sharp end) in about 3–5 minutes. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.4.png‎|300px|left]]
<b>Figure 1.4: Microelectrode filling.</b> The photo should give you an idea of how to fill the microelectrode. First, a small aliquot of 3 M KCl is placed at the back end of the micropipette. Capillary action will cause the KCl solution to fill the tip. Then, the micropipette is backfilled to ensure electrical connectivity.
<br clear=right>
</p>

</td></table>

<p>While you are waiting for the tip to fill, you can fill the microelectrode holder with the same conductive solution (3 M KCl). After 3–5 minutes, re-insert the fine needle into micropipette, all the way to the shoulder (you should see the solution has filled to the shoulder due to capillary action). Fill the shank completely; avoid any bubbles in the glass tubing.</p>

<p><i><strong>The micropipette tip is easily broken! Because of its small size, you may be unaware that it has been broken until you attempt to use it to impale the cell. If you touch, even lightly, the tip to any object, it will break. Having been warned, please exercise due care.</strong></i></p>

<p>The micropipette is now ready to insert in the holder. Again, assure there are no bubbles in the holder or micropipette. Once inserted, tighten the cap to hold the micropipette firmly.</p>

<p><b>Reference electrode.</b> The electrical network must be connected to ground. This is done using the so-called reference electrode. The half cell (Cl<sup>–</sup> + Ag <-------> AgCl + e<sup>–</sup>) is connected to the solution in the cell specimen holder by a salt-bridge (3 M KCl) immobilized in agar (2% w/v). By using 3 M KCl in the salt-bridge, the salt and half-cells are matched for the microelectrode and the reference electrode. You will be provided with a reference electrode pre-filled with 3 M KCl in agar. The reference electrode is stored in 3 M KCl, it is important to rinse off any residual KCl with the distilled H<sub>2</sub>O squeeze bottle. After the cell is placed in the chamber (see below), place the electrode in the holder next to the specimen chamber and adjust so that the tip is immersed. Connect the holder to the electrometer by connecting the silver wire at the back of the reference electrode to the ground on the electrometer. </p>

<p><i><strong>Do not leave the reference electrode in air, so that it dries out. If it does dry out, electrical connectivity will be lost.</strong></i></p>

<p><b>Cell preparation and use.</b> The internodal cells of Chara australis will be provided for you stored in APW7 (artificial pond water, pH 7.0; APW7 contains the following (in mM): KCl (0.1), CaCl<sub>2</sub> (0.1), MgCl<sub>2</sub> (0.1), NaCl (0.5), Mes buffer (1.0), pH adjusted to 7.0 with NaOH.). A single internode cell must be carefully placed in the cell specimen holder. The cell is laid across three chambers. The impalement and reference electrode are located in the central chamber. The two outer chambers are provided with metal leads to generate the voltage stimulus that triggers the action potential. The three chambers are electrically isolated from each other using petroleum jelly, in which the cell is embedded. Small weights will be provided to hold the cell in the specimen chamber, if required.
</p>

<p><i><strong>The cells must not be left in the air! They will be damaged. Furthermore, the cells must be handled gently. Don’t twist or deform them! Either stress (being left in the air or wounding during handling) will harm your ability to obtain good electrical measurements from the cell.</strong></i></p>

<p>Ensure that the reference electrode is touching the solution (see above). If not, the circuit will be incomplete, resulting in wild swings in the measured voltage.</p>

<p>Insert the microelectrode holder into the headstage of the electrometer. By using the micromanipulator, you should be able to place the tip of the micropipette directly above the cell. It is sometimes easier to do this by eye, rather than using the dissecting microscope. Having positioned it, you are now ready to enter the APW7 solution. When the micropipette tip is in the APW7 solution, you will have a complete circuit. Use the test button on the electrometer to inject a 1 nA current through the micropipette. You should see a voltage deflection of about 1–2 mV, corresponding to a resistance of about 1–2 MΩ. If the tip resistance is very high (> 10 MΩ), it is likely that there is a high resistance somewhere else in your circuit. Common problems include the reference electrode and bubbles in the micropipette holder.</p>

<p>The completed set-up, ready for impalement, is shown in schematic form in Figure 1.5.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:01_Figure_1.5.png‎|300px|left]]
<b>Figure 1.5: Impalement setup.</b> The schematic should give you an idea of how the setup will look in preparation for impalement. Not shown are the positioning clamps for the reference electrode and micromanipulator for the microelectrode. For dual impalements, the second headstage-holder-microelectrode assembly is located to the left.
<br clear=right>
</p>
</td></table>

<p>For measurements of cell potential alone (and action potentials), impalements with a single micropipette are all that is required. <i><font color="gray">Aside: For measurements of cell resistance and capacitance, two impalements would have to be performed. The reason for this is that the micropipette tip resistance is similar in magnitude to the membrane resistance. So, it is difficult to separate out these two resistances ''in series''. Furthermore, the tip resistance of the micropipette may change when the cell is impaled. The tip may ‘micro-break’, or cytoplasm may enter the tip; the first scenario will cause a lower resistance, the second will cause a higher resistance (why?). For these reasons, dual impalements are optimal.</font></i></p>

<p><b>Cell impalement.</b> Use the micromanipulator to bring the micropipette tip to the cell. When you (''gently'') press the tip against the cell wall, you may see a dimple in the wall. With additional movement of the tip against the wall, it will ‘pop’ into the cell. Your visual observation of impalement must be verified by a change in the microelectrode potential, from zero to a negative value (your can expect values of about –150 to –200 mV). Sometimes, the potential is initially about –100 mV, but then slowly becomes more negative. </p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:Pulse_train.JPG|300px|right]]
<b>Figure 1.6: Example of a current pulse train.</b>
The voltage is shown in the upper trace, the current pulse is shown in the lower trace. Note the delayed rise in voltage, due to ‘R•C filtering’. All the information required to calculate the capacitance is shown on the graph. The steady state deflection can be used to determine the resistance. The rise time can be determined, and hence the capacitance.
<br clear=right>
</p>
</td></table>

<p><b>Protoplasmic streaming.</b> Not explicitly a part of this lab exercise, protoplasmic streaming may be observable during your experiments. If it is, you may find it interesting to observe the rate and/or direction of the protoplasmic flow. Alternatively, you will be able to view protoplasmic streaming on the Nikon Optiphot microscope with a X10 objective by placing the internodal cell in a Petri dish containing APW7 solution (ensure the cell is immersed). To view protoplasmic streaming during electrical measurements, zoom the magnification on the dissection microscope head to its maximal level.
</p>

<p><b>Action potentials.</b><ref>Johnson BR, Wyttenbach, R.A., Wayne, R., and R. R. Hoy (2002) Action potentials in a giant algal cell: A comparative approach to mechanisms and evolution of excitability. The Journal of Undergraduate Neuroscience Education 1:A23-A27.</ref> Although action potentials may occur spontaneously, it is usually easier (and requires less patience) to induce them. A common method is to stimulate the cell with a large, transient voltage. The voltage is applied between the two outside chambers of the cell holder.
</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:01_micropipette_etc.png|400px|right]]
<b>Micropipette etc.</b> The micropipettes are filled as described previously.
<br clear=right>
</p>
<p align=justify>[[File:02_Chara_tank.png|145px|right]]
<b>Chara in the tank.</b> The Chara is carefully cut from plants in the aquarium by selecting straight internodes and
cutting the internode cells on either side of the one you selected.
<br clear=right>
</p>
<p align=justify>[[File:03_Chara_dish.png|400px|right]]
<b>Chara in the dish.</b> The internode cell is carefully (<b>very gently</b>) placed in a Petri dish filled with Broyar/Barr media (the nutrient solution used to grow Chara). Be warned that salt leakage from the cut internode cells can elevate potassium ion concentrations in the extracellular solution. This can cause depolarized potentials in subsequent measurements. Flushing the dish with fresh Broyer/Barr media should minimize this potential problem.
<br clear=right>
</p>
<p align=justify>[[File:04_Chara_chamber.png|400px|right]]
<b>Chara in the chamber.</b> The internode cell is carefully (<b>very gently</b>) placed in the chamber. The internode cell should fit in the two slots on either side of the central chamber. The slots are filled with vaseline to create an electrical barrier (see below). Note that the internode cell must be immersed in the media used for measurements.
<br clear=right>
</p>
<p align=justify>[[File:05_Chara_vaseline.png|400px|right]]
<b>Vaseline in the slots.</b> The vaseline 'dams are shown in greater detail. <b>Be gentle!</b> Damage to the internode cell can make it quite difficult to measure action potentials.
<br clear=right>
</p>
<p align=justify>[[File:06_Chara_cage.png|400px|right]]
<b>Chara in the Faraday cage.</b> The chamber can be mounted in the Faraday cage, as shown. Don't forget to connect the stimulator leads to the chamber.
<br clear=right>
</p>
<p align=justify>[[File:07_Chara_impaled.png|400px|right]]
<b>Chara is impaled.</b> Now, mount the microelectrode and impale the cell! Once impaled, the dissecting microscope should be zoomed to maximal magnification, so that you will be able to observe the cytoplasmic streaming.
<br clear=right>
</p>
</table>

<p>Once you obtained a stable potential, you can stimulate the cell to induce an action potential. This is done by passing a large current between the two outer chambers using the pushbutton and 9 Volt battery. Press and release the button! Excessive, long-duration current will affect the cell, and obscure your ability to observe the action potential. Upon induction of an action potential, cytoplasmic streaming should cease. This is best documented by measuring the rate of cytoplasmic flow <i>before</i> you induce the action potential, <i>versus</i> after induction. Once an action potential has fired, the cell will enter a refractory period, so be patient if you want to stimulate the cell again. The refractory period will vary, but generally is several minutes. </p>

<p><b>Lab exercise write-up.</b> The details of your write-up are in the hands of the lab coordinator/lecturer. Of especial interest would be the construction of equivalent circuits and their analysis. The schematic in Figure 1.1 is an excellent starting point. What is the effect of the resistance of the solution? Length of the cell (that is, cable properties)? Resistance of the cytoplasm? How do these affect your measurements? You can even test for the effect of the resistance of the APW solution, by placing the microelectrode —in solution— near and far away from the reference electrode. </p>

<p>Resistance and capacitance, when normalized to membrane area, are usually properties of the bilayer membrane, and therefore should be similar for all organisms. Electrical potentials are surprisingly variable, but almost invariably negative-inside. They depend in part upon the concentrations of permeant ions inside and outside of the cell (per calculations using the Goldman-Hodgkin-Katz equation), and on the presence/activity of electrogenic pumps. Chara australis is normally described as existing in either a “K-state” (the potassium conductance dominates, E<sub>m</sub> of about –150 mV) or a “pump-state” (where the H<sup>+</sup> pump creates an electrogenic component, E<sub>m</sub> of about -200 mV).</p>

<p><i><font color="gray">Aside: Because of the additional challenges of performing dual impalements, it is advisable to practice single impalements first. Then, as you become more adept, take on the additional challenge of impaling the cell with two electrodes. The second impalement can be performed similarly, 1 to 2 cm from the first impalement. <b>Resistance measurements.</b> The electrode test button on the electrometer is limited to 1 nA (positive) currents. To obtain a more complete measurement of cell resistance, the ‘stimulus input’ on the electrometer is used. A command pulse with a magnitude and frequency that you select is connected to the stimulus input (Figure 1.6). To measure the capacitance, the pulse should be square (so that you can measure the rise time of the voltage deflection in response to the current injection. Be careful about the magnitude you select: high amperages will damage the cell, low amperages will result in very small voltage deflections. The resistance is calculated from the known current injection (measured at the current monitor output of the electrometer) and the steady state voltage deflection. Note that current is injected through one micropipette, while the voltage deflection is measured at the other micropipette. Multiple measurements are advised, as is using different current magnitudes, so that you can construct a current versus voltage graph.
<b>Capacitance measurements.</b> Using the same current pulse protocol, you can determine the rise time of the voltage deflection. It’s possible to digitize the data and fit it to an exponential function. More commonly, the time required for the voltage to deflect 63% of the final steady state deflection is used. That time, commonly denoted the rise time τ, is equal to R•C (resistance times capacitance) simplifying the calculation greatly. Again, multiple measurements are advised. There is an important control: The resistance of the solution is fairly high due to the low concentration of ions. Thus, even when the microelectrodes are out of the cell, in solution, there will be a voltage deflection, smaller in magnitude than when the microelectrodes are impaled into the cell. To determine the resistance and the capacitance of the cell, the curves must be subtracted (curve<sub>impaled</sub> — curve<sub>external</sub>).</font></i></p>

<h1> The Natural History of ''Chara''<ref> Roger R. Lew, Professor of Biology, York University, Toronto Canada 10 july 2010</ref> </h1>
<p>Although the experimental exercise is focused on direct measurements of the electrical properties of internodal cells of ''Chara'', it seems very appropriate to introduce students to some of the Biophysical Explorations that have been done using ''Chara''. It has been used for about 100 years, and will continue to be used for the foreseeable future.</p>

<h2>Ecology</h2>
<p>An algal species, ''Chara'' is a genus in the botanical family Characeae, which also includes another favorite model organism for biophysicists: ''Nitella''. It is common in freshwater lakes and ponds in Ontario (McCombie and Wile, 1971). It tends to be found in water that is relatively nutrient-rich (but not excessively) based on measurements of the water conductivity (''Chara'' grows well when the conductance is 220–300 µSiemens cm–2) and ‘transparent’ (non-turbid) based on Secchi disk measurements (a common technique for determining how far light travels through the water column, by lowering a marked disk into the water and determining the depth at which the markings are no longer visible) (''Chara'' prefers transparencies of 2.5–6 m).</p>

<h2>Evolution</h2>
<p>The family Characeae is part of an ‘order’ known as Charales, which in turn is part of an algal ‘meta-group’: Chlorophytes (Green Algae). The first fossils that bear strong resemblance to extant Charales are reported from the Silurian period, about 450 million years ago (McCourt et al., 2004). And in fact, much recent research using molecular phylogeny techniques that compare sequence similarities between extant species suggests that the Charales is the phylogenetic group from which land plants arose. A recent molecular reconstruction is shown in Figure 2.1, from McCourt et al. (2004)<ref>McCourt, R.M., Delwiche, C.F. and K.G. Karol (2004) Charophyte algae and land plant origins. TRENDS in Ecology and Evolution 19:661–666.</ref>.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.1.PNG‎|500px|right]]
<b>Figure 2.1: Molecular phylogenetic reconstruction of evolutionary divergences from which land plants evolved (McCourt et al., 2004)</b>
The figure outlines relatedness based on the sequences of a number of genes. Associated characteristics/traits of each group are shown to the left. The traits are those known to occur in land plants. So, they serve as an alternative way to identify the evolution of groups that from which land plants eventually diverged.
<br clear=right>
</p>
</td></table>

<p>
Is this important? Basically, yes. The ‘invasion’ of land by biological organisms was a relatively late event in geological time. It opened the doors to a remarkable diversification of biological species, especially in the major phylogenetic groupings of insects (invertebrates), vertebrates and land plants (especially flowering plants). This was a time when Earth evolved into a form that would be somewhat familiar to us.</p>

<p>You may be surprised to learn that the discovery of Chara-like fossils and their identification relies upon the application of biophysical techniques in a very big way. To determine the structure of a fossilized organism is not easy. It’s rock, after all, and you can’t peel off the layers very easily. For this reason, Feist et al. (2005)<ref>Feist, M., Liu, J. and P. Tafforeau (2005) New insights into Paleozoic charophyte morphology and phylogeny. American Journal of Botany 92:1152–1160.</ref> used “high resolution x-ray synchrotron microtomography”. The fossil samples were irradiated with the very bright monochromatic x-ray beam at the European Synchrotron Radiation Facility while being rotated. From the digital images captured as the sample was rotated, it was possible to reconstruct the three-dimensional ''internal'' structure of the fossil. One example of a reconstruction from their paper is shown in Figure 2.2.</p>

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.2.PNG‎|200px|right]]
<b>Figure 2.2: An example of microtomography of a Charales fossil using a synchrotron x-ray light source (Feist et al., 2005)</b>
The scale bar 150 µm. Thus very small structures can be observed. The value of synchrotron x-ray microtomography is to elucidate very clearly structures that can only be speculated about when looking at the surface of the fossil, or attempting to reconstruct the three-dimensional structure from thin sections cut with a fine diamond blade.
<br clear=right>
</p>
</td></table>

<h2>Membrane Permeability</h2>
<p>[[File:02_Figure_2.2A.PNG‎|300px|right]]Moving to more recent times, our understanding the nature of the plasma membrane in biological cells relied upon measurements that were performed with Characean algae, primarily the work of Collander (1954). He determined the permeability of ''Nitella'' cells to a variety of solutes of varying molecular weight and hydrophobicity (as measured by partitioning between olive oil and water). Synopses of his results have been published extensively in textbooks ever since, because his results were most easily interpreted by proposing that the membrane was lipoidal in nature. We now know that the composition of membranes is mostly phospholipids. These create a bilayer structure with a hydrophobic interior and hydrophilic outer surface. Such a membrane structure naturally lends itself to the creation of an electrical element in cellular function, since the hydrophobic interior of the bilayer acts as an electric insulator.</p>

<h2>Electrical Measurements</h2>
Electrical measurements have been performed on Characean internodal cells for probably 100 years. These measurements include action potentials, although due care should be exercised in nomenclature. Stimuli of various types do induce transient changes in the potential that are similar in nature to those induced in animal cells and are commonly described as action potentials. However, propagation of the electrical signal is not a consistent trait of action potentials in plants (including ''Chara''). Figure 2.3 shows an example of action-potential-like responses in ''Nitella'', from the work of Karl Umrath, who also explored the nature of ''propagated'' action potentials in the sensitive plant Mimosa.

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.3.PNG‎|500px|right]]
<b>Figure 2.3: An example of an electrical trace from the Characean alga Nitella (Umrath, 1933)<ref>Umrath, K. (1933) Der erregungsvorgang bei Nitella mucronata. Protoplasma 17:258–300.</ref></b>
I can’t offer any explanation of the trace, because the original paper is in German. The first transient upward (depolarizing?) peak (A) is reminiscent of an action potential. The second peak (B) looks like a voltage response to current injection.
<br clear=right>
</p>
</td></table>

<p>The scope of research that has explored the electrophysiology of Characean algae is immense. One important research theme was the nature of electrogenicity; that is, the cause of the highly negative inside potential of the internodal cell. The central role of an ATP-utilizing H+ pump was proposed from research using Characean algae (Kitasato, 1968<ref>Kitasato, H. (1968) The influence of H+ on the membrane potential and ion fluxes of Nitella. Journal of General Physiology. 52:60–87. </ref>; Spanswick, 1972<ref>Spanswick, R.M. (1972) Evidence for an electrogenic ion pump in Nitella translucens. I. The effects of pH, K+, Na+, light and temperature on the membrane potential and resistance. Biochimica et Biophysica Acta 288:73–89.</ref>), and extended to fungi and higher plants. The electrogenic H+ pump plays a role as central as that of the very similar Na+ K+ pump of animal cells. It is important for Biophysics students to be aware of the concept: Choose the right organism for a particular biological problem. In the case of the Characean algae, their large size made it possible to address the biological question of electrogenicity very directly. That is, the experimentalist could cut open the cell, remove the central vacuole, and then fill the cell with any desired solution. From such internal perfusion experiments, the ATP-dependence of electrogenicity was demonstrated directly (Shimmen and Tazawa, 1977<ref>Shimmen, T. and M. Tazawa (1977) Control of membrane potential and excitability of Chara cells with ATP and Mg2+. Journal of Membrane Biology 37:167–192.</ref>). An example of a perfusion setup is shown in Figure 2.4.</p>

<table width=800 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.4.PNG‎|500px|right]]
<b>Figure 2.4: Internal perfusion of an internodal Chara cell (Tazawa and Kishimoto, 1968)<ref>Tazawa M. and U. Kishimoto (1968) Cessation of cytoplasmic streaming of Chara internodes during action potential. Plant & Cell Physiology. 9:361–368</ref></b> The cut ends are submersed in an artificial cell sap in the wells A and B. Not only can this technique be used for model organisms like ''Chara'' but also for animal cells of similar large size. Thus, internal perfusion was used to good advantage in initial studies of the action potential of the squid giant neuron, a system also used to unravel mechanisms of pH regulation.
<br clear=right>
</p>
</td></table>

<h2>Molecular Motors</h2>
Protoplasmic streaming is something you should be able to observe during your experimental exercise. The first experiments on protoplasmic streaming in ''Chara'' date back to the early-1800’s, when Dutrochet performed surgical ligations of the internodal cells (''op cit.'' Kamiya, 1986<ref>Kamiya, N. (1986) Cytoplasmic streaming in giant algal cells: A historical survey of experimental approaches. Botanical Magazine (Tokyo) 99:441–467.</ref>). Kamiya (1986) describes the many other micromanipulations that were used to elucidate the biophysical properties of protoplasmic streaming. What makes this phenomenon even more fascinating is the unabated interest in protoplasmic streaming in ''Chara'' that continues to this day. On the one hand, it is now clear that ''Chara'' has the fastest known myosin molecular motor known (this is the motive force for protoplasmic streaming) (Higashi-Fujime et al., 1995<ref>Higashi-Fujime, S., Ishikawa, R., Iwasawa, H., Kagami, O., Kurimoto. E., Kohama, K. and T. Hozumi (1995) The fastest actin-based motor protein from the green algae, ''Chara'', and its distinct mode of interaction with actin. FEBS Letters 375:151–154.</ref>; Higashi-Fujime and Nakamura, 2007<ref>Higashi-Fujime, S. and A. Nakamura (2007) Cell and molecular biology of the fastest myosins. International Review of Cell and Molecular Biology 276:301–347.</ref>; Ito et al., 2009<ref>Ito, K., Yamaguchi, Y., Yanase, K., Ichikawa, Y. and K. Yamamoto (2009) Unique charge distribution in surface loops confers high velocity on the fast motor protein Chara myosin. Proceedings of the National Academy of Sciences (USA) 106:21585–21590.</ref>). The molecular mechanism of the Chara myosin motor was studied by the optical tweezer technique: Kimura et al. (2003)<ref>Kimura, Y., Toyoshima, N., Hirakawa, N., Okamoto, K. and A. Ishijima (2003) A kinetic mechanism for the fast movement of ''Chara'' myosin. Journal of Molecular Biology 328:939–950.</ref> held an actin filament with dual optical tweezers (one at each end) and measured the force as the actin filament was displaced by the step-wise motion of the ''Chara'' myosin motor. One the other hand, protoplasmic streaming offers insight into the interplay between mass flow and diffusion in the context of micro-fluidics (Goldstein et al., 2008<ref>Goldstein, R.E., Tuval, I. and J-W van de Meent (2008) Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Proceedings of the National Academy of Science, USA 105:3663–3667.</ref>), as measured by magnetic resonance velocimetry (van de Meent et al., 2009<ref>Van de Meent, J-W., Sederman, A.J., Gladden, L.F. and R.E. Goldstein (2009) Measurement of cytoplasmic streaming in Chara corallina by magnetic resonance velocimetry. arXiv:0904.2707v1 [physics.bio-ph]</ref>).

<h2>Future Research</h2>
The continued use of Characean algae in biophysical research is certain. What research will be done? That depends upon the development of new technologies, and the pressing need to elucidate fundamental biological research problems. Braun et al. (2007)<ref>Braun M., Foissner, I., Luhring, H., Schubert H. and G. Theil (2007) Characean algae: Still a valid model system system to examine fundamental principles in plants. Progress in Botany 68:193–220.</ref> describe some of the biological research problems that can be addressed using this “model system ''par excellence'' to study basic physiological and cell biological phenomenon in plants”. These include pattern formation, sensing of gravity and growth responses thereof, polarized growth, cytoskeleton dynamics, photosynthesis (especially coordinated metabolic pathways that involve multiple organelles), wound-healing, calcium signaling, and even sex determination. As to new technologies, the use of nmr imaging and even magnetic field measurements using a superconducting quantum interference device magnetometer (Baudenbacher et al., 2005<ref>Baudenbacher F., Fong, L.E., Thiel G., Wacke, M., Jazbinsek V., Holzer, J.R., Stampfl, A. and Z. Trontelj (2005) Intracellular axial current in Chara corallina reflects the altered kinetics of ions in cytoplasm under the influence of light. Biophysical Journal 88:690–697.</ref>) suggest that as new technologies are developed, biophysicist will turn to ''Chara'' as a tried and true model system for validation of new techniques.

{|border="1"
|+<b>''Chara'' species list</b>
|colspan="3"|
Identifying Chara to species often requires careful examination of the sexual organs. Thus, the list below may include species that in fact are not valid. Nevertheless, it serves as an indicator of the diversity present solely in this one, very famous, genus of the Chlorophyta

|-
|colspan="3"|<b>Sources:</b><ul><li>Encyclopedia of Life (http://www.eol.org/pages/11583)(black)</li>
<li>NCBI (http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=9421)(red)</li>
<li>UniProt (http://www.uniprot.org/taxonomy/13778)(blue)</li></ul>
|-
|Chara australis (corallina)
|Chara halina A. García +
|Chara pouzolsii J. Gay ex A. Braun +
|-
|Chara curtissii
|Chara hispida Linneaus +
|Chara preissii
|-
|Chara denudata Desvaux & Loiseaux +
|Chara hookeri A. Braun +
|Chara pulchella +
|-
|Chara drouetii
|Chara hornemannii Wallman
|Chara rudis (A. Braun) Leonhardi +
|-
|Chara dichopitys +
|Chara horrida Wahlstedt +
|Chara rusbyana M. Howe +
|-
|Chara eboliangensis S. Wang +
|Chara huangii Y. H. Lu +
|Chara sadleri F. Unger +
|-
|Chara ecklonii A. Braun ex Kützing +
|Chara hydropytis +
|Chara sejuncta A. Braun, 1845
|-
|Chara elegans (A. Braun) Robinson
|Chara imperfecta A. Braun +
|Chara setosa Klein ex Willd. +
|-
|Chara excelsa Allen, 1882
|Chara intermedia A. Braun +
|Chara spinescens Fée +
|-
|Chara fibrosa C. Agardh ex Bruzelius +
|Chara leei S. Wang +
|Chara stoechadum Sprengel +
|-
|Chara foetida
|Chara leptopitys A. Braun +
|Chara strigosa
|-
|Chara foliolosa
|Chara longifolia
|Chara stuartiana
|-
|Chara formosa Robinson, 1906
|Chara montagnei A. Braun +
|Chara tomentosa Linneaus +
|-
|Chara fragilifera Durieu +
|Chara muelleri
|Chara vandalurensis
|-
|Chara fragilis Loiseleur-deslongchamps, 1810
|Chara muscosa J. Groves & Bullock-Webster +
|Chara virgata Kützing +
|-
|Chara galioides De Candolle +
|Chara palaeofragilis +
|Chara visianii J. Blazencic & V. Randjelovic +
|-
|Chara globularis Thuiller +
|Chara polyacantha
|Chara vulgaris Linnaeus +
|-
|Chara gymnopitys
|Chara polycarpica Delile +
|Chara wallrothii Ruprecht +
|-
|Chara haitensis
|
|Chara zeylanica Klein ex Willdenow +
|-
|}

<table width=700 border=1 align=center><td>
<p align=justify>[[File:02_Figure_2.5.PNG‎|400px|right]]
<b>''Chara'' Life Cycle</b> Various structures of the ''Chara'' life cycle are shown (from Lee, 1980)<ref>Lee, R.E. (1980) Phycology. Cambridge University Press. pp 441–445.</ref>. The nucule is the female organ, the globule is the male organ. The eggs are fertilized by a motile sperm cell (antherozoid). The sperm cells swim with a twisting motion (almost a Drunkard’s walk) as they search for the egg cells. Once fertilized, the zygospore (diploid) may remain dormant for extended periods of time (up to 40 years based on recovery of germinating material from old lake sediments). Once released from dormancy (which requires light), the first division is meiotic, so that the vegetative structures are haploid. Rhizoids grow into the pond sediment. Once anchored, the filamentous internode and whorl cells grow upward towards the water surface. The internode cells are multinucleate.
<br clear=right>
</p>
</td></table>

<h2>References</h2>

<references/>

<h1> Appendix One: micropipette physics </h1>
[[File:03_Appendix_1.png|700px|center]]
[[File:03_Appendix_2.png|700px|center]]

<h1> Appendix Two: electrometer circuitry </h1>

<p>Although it is outside the scope of the lab exercise, it is often helpful to know what is happening ‘inside the box’. So, an example of an electrical schematic for an electrometer is shown below. The electronics are designed to work under high impedance conditions. That is, because of the high resistance of the microelectrode, the input impedance of the electrometer must be at least 10<sup>3</sup> higher (to avoid underestimating the voltage because of voltage dividing). Normally, the ‘heart’ of the electrometer is the electrical circuitry in the headstage of the electrometer. This is where the low noise voltage follower is housed, as near as possible to the cell being measured to minimize the extent of electrical noise pick-up. The headstage is connected to the electrometer with a shielded cable. Thus, in a ‘normal electrometer’, the circuitry shown below is housed in two separate enclosures.</p>

<table width=900 border=1 align=center><td>
<p align=justify>[[File:Electrometer_circuitry.JPG‎|500px|right]]
<b>Figure 4.1: Electrical schematic of a typical high input impedance electrometer. </b> This example is not completely equivalent to the circuit in the electrometers you will be using. It is a simplified circuit from a book by Paul Horowitz and Winfield Hill (1989) The Art of Electronics. Cambridge University Press. pp. 1013–1015. Low-noise FET (field effect transistor) operational amplifiers (IC1 and IC2) that have low input current noise (to minimize voltage offsets caused by the high resistance of the cell (>10<sup>6</sup> Ohms) and input impedance of the electrometer (>10<sup>11</sup> Ohm). The two operational amplifiers are configured as an instrumentation amplifier, in which the voltage is measured as the difference between cell voltage and ground. This is a very effective way to remove extraneous electrical noise from the measurement, known as common mode rejection (CMR). The FETs are usually carefully paired to match voltage drift. The rest of the circuit includes active compensation of capacitance and a reference voltage to provide greater measurement accuracy.
<br clear=right>
</p>
</td></table>

<p>To further minimize the problem of electrical noise pick-up because of the high impedance of the microelectrode, it is normal to ‘shield’ the cell specimen holder from electromagnetic noise (fluorescent lamps and computer monitors are common sources of electrical noise). The shield —usually an enclosing grounded box of conductive metal— is called a Faraday cage. You should be able to observe the problem of extraneous noise yourself. With the microelectrode and reference electrodes in place, stick your finger near the cell specimen chamber while pointing your other arm at the overhead fluorescent lamps. You should see the appearance of noise on the DC output of the electrometer, probably in the frequency range of 60 Hz. </p>