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Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy

2011-01-17T21:03:26Z

Sbilling: 31 revisions

<p>In this experiment, you will be using studying a fragment of a protein called CASKIN. It is involved in neuronal signaling. When a neuron is stimulated, chemical messengers are transferred between two neurons at a specialized cell-cell junction called a synapse. CASKIN is a scaffolding molecule found at the pre-synaptic side of the connection. A scaffolding molecule, much like its name, helps assemble other protein together in a certain part of the cell.</p>

<ul><li>In the box below, draw a schematic of a synapse and the approximate location where CASKIN.</li></ul>

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<p>CASKIN is composed of many protein domains, each with its own function. Not suprisingly, since CASKIN’s job in the cell is to bring together other proteins, it is composed exclusively of domains that promote protein-protein interactions.</p>
<p> Below is a schematic of human CASKIN (1365 amino acids). This schematic was retrieved from the UNIPROT website (http://www.uniprot.org/uniprot/Q8WXD9), a respository of information on thousands
of proteins. We will be studying the SH3 domain of CASKIN, found between amino acids 281 and 347 in the protein.
[[File:Nmr_figure1.jpg‎|500px|center]]
Members of the Donaldson laboratory used a technique called PCR to amplify the gene fragments encoding the SH3 domain, the first SAM domain, the second SAM domain, and the two SAM domains together. These fragments were inserted in to an expression vector, a circular piece of DNA or plasmid, that can be introduced into the common laboratory bacterium, Escherichia coli. Upon addition of a special kind of sugar into the culture which the bacteria cannot metabolize, the bacteria will begin to produce the human protein that we have programmed into them.</p>

<p>The cells were harvested and then cracked open. From the thousands of proteins in the bacterium the CASKIN SH3 domain was purfied in about two hours because it was tagged with a special sequence of six histidines that no other native bacterial protein had. Six histidines bind nickel very well so we used a special type of chromatography when nickel ions are exposed on a resin.</p>
<p>
The protein was further purified by a technique called gel filtration that separated molecules according to their size. At this point the protein was extremely pure. It was then concentrated to about 10 mg /mL which doesn’t sound like much but it is !</p>

<ul><li>From the NMR laboratory tour that you participated in, fill in the vital statistics of the CASKIN SH3 domain NMR sample:</li></ul>
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<tr><td>
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</td>
</table>
</td>
<td>length of NMR tube</td>
</tr>
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</td>
</table>
</td>
<td>width of NMR tube</td>
</tr>
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</td>
</table>
</td>
<td>volume range of NMR sample (mL)</td>
</tr>
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</td>
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</td>
<td>ideal concentration of a protein [mM]</td>
</tr>
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</td>
</table>
</td>
<td>temperature range of experiments</td>
</tr>
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</td>
</table>
</td>
<td>temperature of the magnet (K)</td>
</tr>
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</td>
</table>
</td>
<td>field strength of the magnet (Tesla)</td>
</tr>
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</td>
</table>
</td>
<td>precession frequency of a 1H nucleus</td>
</tr>
<tr><td>
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</td>
</table>
</td>
<td>precession frequency of a 15N nucleus</td>
</tr>
</table>

<p>The protein that was used to acquire the NMR data was uniformly isotropically labeled with 15N. To achieve this, the bacteria were grown in a medium containing 15N-ammonium chloride as their only source of nitrogen.</p>

<hr>
<ul><li>Why is 15N useful for NMR spectroscopy and not 14N, the natural abundance isotope ?</li></ul>
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<p>One of the first NMR experiments that is acquired on a protein sample is called a 2D 15N-edited HSQC. This experiment consists of a pulse sequence that selects only for 1H bound by one bond to a 15N. All other bonds such as 1H-12C and 1H-13C are not observed. An HSQC experiment is an extremely useful tool to gain and immediate understanding of the protein structure, it’s like a fingerprint of sorts.
Here is the basic chemical formula of a protein. As you can see, there is an amide NH in the protein backbone. Thus, you would expect to see one NH resonance (or spot/peak) per amino acid. The CASKIN SH3 domain that we will be studying has the following sequence:</p>
[[File:Nmr_figure2.jpg‎|500px|center]]
<p>The quaternary amino group (NH3+) at the beginning of a protein is not visible by NMR methods. The 1H exchange with protons from water very quickly.</p>
<p>Some amino acids have NH in their side chains and other amino acids do not have any NH at all. Here is the sequence of the CASKIN SH3 domain exactly how it was made for the NMR experiments.</p>

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<p align=justify>[[File:Nmr3_sequence.png|400px|center]]
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</p>
</td></table>

<ul><li>Consulting a table of amino acids, how many NH resonances would you expect to see in a 2D spectrum of the CASKIN SH3 domain ?</li></ul>
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<p>Here is an HSQC spectrum of the CASKIN SH3 domain. The number of resonances is probably lower than what you expected from your calculation. This is a portion of a large NMR spectrum in the chemical shift range that you would expect to find 1H and 15N in a protein. The center of the true NMR spectrum is 4.7 ppm (though all amide NH are found downfield, or at larger values than that). The center of the true NMR spectrum is 120 ppm for 15N or exactly what is shown there.</p>
<p>Using a series of 3D NMR experiments, I assigned (or identified) the specific amino acid in the SH3 domain that contributed a given resonance. This procedure required about one week of data collection from the NMR spectrometer and about four hours of manual inspection. There are computer programs available now that can assign the spectrum in a matter of minutes, but the dataset has to be excellent (high signal-to-noise, no artifacts). For many proteins, that is not possible, although the programs are getting better with handling imperfect datasets.</p>

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<p align=justify>[[File:Nmr4_plot.png|700px|center]]
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</p>
</td></table>

<p>A special filtering technique was used to remove all of the NH2 amino groups from the spectrum so there are only NH’s visible. Note that all of the peaks are spread out. Inituitively, you might think that the spectrum might be a big blob of peaks in the middle since they are, in essence, all NHs and chemically the same. That’s true, but in a protein, each amide NH belongs to a different amino acid which creates some electronic differences which are then manifested as magnetic differences. The biggest contributing factor, though, is that the protein is structured. And when an NH participates in an hydrogen bonds (in an alpha helix or beta sheet) or is near an aromatic group (due to the configuration of their electrons, they are like little magnets themselves), their local environment becomes different. For some small chemical compounds, there are programs that can predict very well where a certain 1H resonance should be found. However, these computing methods rely on quantum mechanics principles and the number of atoms to deal with in a proteins makes the solution impossible to calculate.</p>
<p>The great thing about chemical shifts is that they are very sensitive. Changes the chemical enviroment (and structure) of the protein will be observed as changes in chemical shift. For example,
<ul>
<li>pH</li>
<li>temperature</li>
<li>salt concentration</li>
<li>protein modifications such as phosphorylation</li>
</ul>
will all change the chemical shifts of a protein.</p>
<p>Chemists studying an enzymes mechanism like to perform chemical shift perturbation Amino acid sidechains that are ionizable in the active site (such as Asp, Glu, His, Arg, Lys) and catalytically important will often display different individual pK’s that are distinct from non-catalytic side chains.</p>
<p>Pharmaceutical researchers use chemical shifts in a method SAR-by-NMR (Structure Activity Relationship by NMR). Instead of altering chemical conditions, they add a drug to the solution. If the drug binds somewhere on the protein, the chemical shifts in the vicinity of the drug will generally changes as the protein changes its structure to bind the drug. If the chemists know the identity of the changes, they can map those changes onto a structure of the protein and identify where the drug is binding. So more clever approaches involve binding a different drug somewhere nearby and then tethering the two identified drug neighbours into a more potent superdrug that is more than the sum of its parts.</p>
<p>SH3 domains are well known for their ability to bind short 7-9 amino acid long peptides that tend to be rich in proline. Take a look at Dr. Tony Pawson’s excellent website to learn more:</p>

http://pawsonlab.mshri.on.ca/index.php?option=com_content&task=view&id=179&Itemid=64

<table width=600 align=center><td>
<p align=justify>[[File:Nmr5.png|400px|left]]
Here is an SH3 domain in ribbon format that shows its series of beta strands coloured
green. A peptide in orange binds a set of grooves in the surface of the protein. The peptide is special in that it adopts an left-handed helix. Generally, only proline-rich sequences can form a left-handed over the more common right handed helix
<br clear=right>
</p></table>

<p>Your next objective is to interpret some binding data on the following pages. To the CASKIN SH3 domain, I added a hypothetical ligand for it. As a result some of the amide chemical shifts changed. You first task is to compare the control spectrum (free SH3 domain) and the experiment spectrum (SH3 domain plus ligand) and identify the changes and their magnitude.</p>
<p>Although I would do it by computer, you can do it by overlaying the two spectra and first identifying the major movers. You can get the magnitude of the changes simply by using a ruler and measuring the distance. A distance in mm doesn’t need to be converted back into ppm because you are just scaling by a constant value anyways.</p>

<ul><li>List the amides that moved the most below, along with the magnitude of the change:</li></ul>

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<ul><li>Analyse the labeled reference spectrum provided on the earlier pages. Which amide resonances are missing ? Notice that some peaks are weaker than others. It turns out that sites on the protein that are experience additional dynamics in the range of miili- to microseconds tend to have peaks broadened out. The volume is the same - it’s just the peak is really wide. List the missing and weak resonances below:</li></ul>

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<p align=justify>[[File:Nmr6.png|800px|center]]
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</p>
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<table width=400 align=center><td>
<p align=justify>[[File:Nmr7.png|800px|center]]
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</p>
</td></table>

<p> Now that you have your chemical shift perturbation data, you can map it onto the surface of the CASKIN SH3 domain structure.
Download the structure as a pyMOL session file from</p>

http://farq047c.biol.yorku.ca/biophysics/caskinsh3.pse

<p>Download the pyMOL molecular graphics program from </p>

<p>Mac:</p> http://www.pymol.org/rel/099/macpymol-0_99rc6.tar.gz
<p>Windows:</p> http://www.pymol.org/rel/099/pymol-0_99rc6-bin-win32.zip

<p>If everything worked with pyMOL and the file, you should have a screen like the one below:</p>

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<p align=justify>[[File:Nmr8.png|700px|center]]
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</p>
</td></table>

<p>You can make your biggest movers, slight movers and weak peaks by making selections in the scripting language of the program. For example, to make a selection of amino acids 19, 35 and 52, I could creating a special “object” called “an-example” by typing according to the syntax below. Try it for your various movers and such:</p>

<table width=600 align=center><td>
<p align=justify>[[File:Nmr9.png|700px|center]]
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</p>
</td></table>

<p>You can colour and annotate using the buttons to the right of the selection. “S” is for Show. “H” is for Hide. “L” is for Label and “C” is for colour.</p>
<p>Here I’ve clicked on the “S” and “spheres” selections in the “an-example” entry to highlight those amino acids in space filling format.</p>

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<p align=justify>[[File:Nmr10.png|700px|center]]
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</p>
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<p>Using the “C” button, you can choose a colour for them. I chose green.</p>

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<p align=justify>[[File:Nmr11.png|700px|center]]
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</p>
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<p>In this next example, I went to the “S” button in “top10-1”, that’s the name of the original molecule, and selected “spheres”. Now I can see where the highlighted amino acids are on the surface.</p>
<p>You can save what you are doing at any time by going to the main menu under File and choosing “save session as”</p>

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<p align=justify>[[File:Nmr12.png|700px|center]]
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</p>
</td></table>

<p>This is part where you are now on your own ! With your data and pyMOL, figure out where the binding surface is on the molecule. Also where are missing amino acids located ? At the loops, at the ends of the molecule ? Somewhere else?</p>

<p>You can save a nice picture by taking a screen-shot or by using the “png” command.</p>

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<p align=justify>[[File:Nmr13.png|700px|center]]
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</p>
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<p>Consult the comprehensive “pyMOL wiki” for more commands and tips</p>

http://www.pymolwiki.org/index.php/Main_Page

<p>Attach your output to the end of this laboratory report along with any additional information and insights. Good luck !</p>

Main Page/BPHS 4090/ElectroPhysiology of Chara revised

2011-01-17T21:03:24Z

Sbilling: 15 revisions

<h1>Overview</h1>

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<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.
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</p></table>

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<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.
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<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]]
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</p></table>

<h1>Lab Exercise<ref>By Roger R. Lew, Professor of Biology, York University, Toronto Canada. 06 september 2010.</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) Resistance and capacitance measurements (dual microelectrode impalements).
</p>
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<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.
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<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>

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<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).
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<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>

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<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.
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</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. Small weights will be provided to hold the cell in the specimen chamber.
</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>

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<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.
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</p>
</td></table>

<p>For measurements of cell potential alone, impalements with a single micropipette are all that is required. For measurements of cell resistance and capacitance, two impalements must 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.</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>

<p><i><strong>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.</strong></i></p>

<p>The second impalement can be performed similarly, 1 to 2 cm from the first impalement.</p>

<p><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. </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>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.</p>
<p>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>).</p>

<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).
</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>

<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.
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</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 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/Lysozyme Crystallization

2011-01-17T21:03:20Z

Sbilling: 20 revisions

<h1>Introduction</h1>

<h2>Three-dimensional Protein Structures</h2>
<h2>X-Ray Crystallography</h2>
<h2>What is a Protein Crystal? </h2>
<h2> Suggested Reading:</h2>

<p><b>Introduction to Macromolecular X-Ray Crystallography</b> by Esko Oksanen and
Adrian Goldman (Chapter 10 in Comprehensive Natural Products II Chemistry and
Biology, <i>Editors-in-Chief: Lew Mander and Hung-Wen (Ben) Liu</i> ISBN: 978-0-08-
045382-8).</p>

<p>Alexander McPherson. <b>Introduction to protein crystallization.</b> Methods, 34, 254-265.</p>

<p><b>Protein X-Ray Crystallography Methods</b> by Roger S. Rowlett, Department of
Chemistry, Colgate University</p>

<h2>Websites</h2>
<ul>
<li>http://www.ruppweb.org/level1/new_tutorials_page.htm</li>
<li>http://www.sdsc.edu/Xtal/edu.index.html</li>
<li>http://www.hamptonresearch.com</li>
<li>http://www.pxuniverse.com/</li></ul>

<h1>Experimental</h1>
<p>You will be provided with the pure protein (lysozyme) that you will be crystallizing. You
will also be provided with the crystallization solution. In real life you perform
crystallization trials with your homogeneously pure protein and examine your plates
looking for protein crystals.</p>

<h2>Crystallization Experiment I</h2>
<h3>Stock Solutions</h3>
<h3>Protein</h3>
<p>Lysozyme at 25, 50, 75 and 100 mg/ml dissolved in 0.02 M sodium acetate pH 4.6</p>
<h3>Lysozyme Crystallization Solution</h3>
<ul>
<li>0.6 M sodium chloride</li>
<li>0.1 M sodium acetate pH 4.6</li>
<li>25% Glycerol</li></ul>
<h3>Procedure</h3>
<table width=280 align=left ><td>
<p align=justify>[[File:Fig1_crystallization_plate.png|280px|center]]
<b>Figure 1:</b> Crystallization Plate
<br clear=right>
</p>
</td></table>
<br style="clear: both" />
<ol>
<li>Fill reservoirs with 500 μl of lysozyme crystallization solution.</li>
<li>Add 1 μl of the crystallization solution to 1 μl of lysozyme on the cover.</li>
<table width=200 align=left ><td>
<p align=justify>[[File:Figure2.png|200px|center]]
<b>Figure 2</b>
<br clear=right>
</p>
</td></table>
<br style="clear: both" />
<li>Repeat for the remaining lysozyme concentrations. You should have 4 drops on
each cover. Keep in mind where you place the different lysozyme concentrations by
referring to the notch.</li>
<li>Screw cover on reservoir.</li>
<li>Repeat for the remaining reservoirs in your row.</li>
<li>Incubate at room temperature.</li>
<li>Observe and document conditions in drops with microscope immediately after setup
and 1, 2 and 3 weeks later.</li>
</ol>

<p>Describe the results! Do you see any trends? How many crystals do you see per drop?
Which drop contains the largest crystals? Which drop contains the most crystals?</p>

Describe Vapor Diffusion as it pertains to crystal growth.
<table width=400 align=left ><td>
<p align=justify>[[File:Fig3_vapour_diffusion.png|400px|center]]
<b>Figure 3:</b> Well setup for crystal growth by the vapour diffusion method
<br clear=right>
</p>
</td></table>
<br style="clear: both" />

<h2>Crystallization Experiment II</h2>
<h3>Stock Solutions</h3>
<h3>Protien</h3>
<p>Lysozyme at 25, 50, 75 and 100 mg/ml dissolved in 0.02 M sodium acetate pH 4.6</p>
<h3>Crystallization Conditions</h3>
<h3>Qiagen JCSG+ Screen</h3>
<p>This crystallization screen contains 96 known solutions. You will use 24 of these
conditions to setup an initial crystallization screen for the 4 different lysozyme
concentrations.</p>
<p>Prepare the crystallization plate as above but remember to use a different crystallization
condition in each well. The crystallization conditions are labeled 1-24. Each group will
set up 6 different conditions in one row.</p>
<p>Observe and document the drops with microscope. Shown below are typical drop
results. Most drops will be either clear (no crystals, no precipitate) or contain precipitate
(protein).</p>
<p>Describe the results! Do you see any trends? How many drops are clear? How many
drops have precipitate? Do any drops contain crystals?</p>

<table width=500 align=left ><td>
<p align=justify>[[File:Figure4_gallery.png|500px|center]]
<br clear=right>
</p>
</td></table>
<br style="clear: both" />

<h2>Protien Crystal Mounting</h2>
<p>Many protein crystals are extremely sensitive to changes in temperature and
surrounding conditions (i.e., presence of original solvent). As well, the protein crystals
5
undergo radiation damage upon exposure to X-rays. Therefore, the crystal must be
flash-cooled to 100K in a stream of liquid nitrogen to prevent damage from the highintensity
x-rays. Cryoprotectants such as glycerol are used to prevent freezing of the
water surrounding the protein crystal which may destroy the protein crystal.</p>

<table width=300 align=left ><td>
<p align=justify>[[File:Fig5_pins.png|300px|center]]
<b>Figure 5:</b> Different pins containing a loop on the top end.
<br clear=right>
</p>
</td></table>
<br style="clear: both" />

<table width=300 align=left><td>
<p align=justify>[[File:Fig6_mount.png|300px|center]]
<b>Figure 6:</b> Using a pin to mount a crystal in a loop.
<br clear=right>
</p>
</td></table>
<br style="clear: both" />

<table width=300 align=left ><td>
<p align=justify>[[File:Fig7_loop.png|300px|center]]
<b>Figure 7:</b> Using a pin to mount a crystal in a loop observed through
the microscope. The size of the crystal dictates the size of the loop. The crystal will be
scooped into the loop and mounted on the goniometer head for diffraction analysis.
<br clear=right>
</p>
</td></table>
<br style="clear: both" />

<p>You will be given an opportunity to mount the lysozyme crystals into the loops during
the lab.</p>

<table width=600 align=left ><td>
<p align=justify>[[File:Fig8_setup.png|600px|center]]
<b>Figure 8:</b> Showing the pin containing the protein crystal mounted on the goniometer head
on the diffractometer.
<br clear=right>
</p>
</td></table>
<br style="clear: both" />

<p>You will test the diffraction potential of your lysozyme crystals using a Rigaku 007HF
Microfocus X-ray machine equipped with a Saturn 944+ CCD.</p>

<p>Describe the results. What is the diffraction resolution of your lysozyme crystals? What
is the space group?</p>

Main Page/BPHS 4090/In-Vivo Spectrocopy

2011-01-17T21:03:19Z

Sbilling: 72 revisions

<h1> Required Components </h1>

<ol>
<li>[[Media:Spec_Spectrometer.JPG|USB Ocean Optics spectrometer]]</li>
<li>[[Media:Spec_Collection_Fiber.JPG|Collection fiber for spectrometer]]</li>
<li>PC with spectrometer software and MS Excel (or equivalent spreadsheet) for data processing</li>
<li>[[Media:Spec_White_Reference.JPG|White reflectance reference target]]</li>
<li>[[Media:Spec_Light_Source.JPG|White LED illumination source]]</li>
<li>[[Media:Spec_Pressure_Cuff.JPG|Blood pressure compression cuff]]</li>
<li>Excel calculation spreadsheets</li>
<li>Stopwatch timer (can use windows clock for this)</li>
<li>A usb stick or some way to take your data with you for analysis</li>
</ol>

<h1>Objective</h1>
<p>To measure the in-vivo oxygenation state of haemoglobin, and calculate the change in oxygenation before, during, and after reactive hyperaemia by analyzing the colour content of light diffusely reflected off of the skin.</p>

<h1>Introduction</h1>
<p>Optical methods of skin analysis are ideal because they can be performed non-invasively, and in real-time. It is quite intriguing that so much information can be discovered from something as simple as launching some photons at an object and analyzing what comes back. In this lab, you will be exploring the use of light as a non-invasive measurement tool to determine the in-vivo oxygenation status of haemoglobin in your blood. These measurements will be made in a non-invasive sense, so as much as you may enjoy slicing up your lab partner to get at their blood, it ain’t gonna happen here! You will, on the other hand, have the opportunity to cut off the blood flow to one of your lab partner(s) limbs, though sadly, this will only be temporary. In this lab you will get familiar with the concept of light propagation in turbid (scattering) media, as well as gain experience with optical spectroscopy methods. High resolution spectral information can be analyzed to allow semi-quantitative and fully quantitative analysis of biological materials, and is a very powerful technique, useful for a variety of biophysical applications.</p>
<table width=280 align=left ><td>
<p align=justify>[[File:Absorption_spectrum_of_melanin.JPG|240px|border|center]]
<b>Figure 1</b> - Absorption spectrum typical for melanin.
<br clear=right>
</p>
</td></table>
<p>Determination of physiologically relevant parameters in a quick, reliable and repeatable fashion is of paramount importance in healthcare and biological research. The optical properties of human skin have been the subject of numerous investigations over the years, and two of the most relevant parameters to measure are the haemoglobin (Hb) oxygenation state and melanin content. Hb and melanin are the two major cutaneous chromophores within human skin, which means that their concentrations are essentially responsible for the colour of your skin. Upon exposure to ultraviolet (UV) light, melatinocytes increase their production of melanin within the skin, we know this process by its more common name, a suntan. The absorption spectrum of melanin is shown in figure 1, and is almost linear over the visible spectrum. It is best measured in the spectral rang above 600 nm, as it is the main source of light absorption in the skin at this wavelength and beyond.</p>

<p> The main target we are after in this lab is the oxygenation state of Hb. Hb is the iron-containing protein attached to red blood cells, and aside from giving blood its red colour, it is responsible for transporting oxygen from the lungs to the rest of the body. The mechanism of oxygen binding in Hb is due to a single iron atom, contained within the protein structure of Hb, and just below a porphyrin ring. A 2D representation of the ring and iron is shown in Figure 2. This structure serves to trap an oxygen molecule and hold it for transport around the body. A typical Hb molecule consists of four of these binding sites surrounded by a protein matrix, and the overall structure of the Hb molecule changes when carrying oxygen. This structural change results in a change in the absorption spectrum of haemoglobin in the 400-600 nm spectral range (Figure 3). </p>
<br style="clear: both" />
<table width=280 align=left><td>
<p align=justify>[[File:Haeme_ring.jpg|240px|border|center]]
<b>Figure 2</b> - Schematic representation of the haeme porphyrin ring in Hb.
<br clear=right>
</p>
</td></table>
<table width=280 align=left><td>
<p align=justify>[[File:Absorption_spectrum_of_Hb.jpg|240px|border|center]]
<b>Figure 3</b> - Absorption spectra of oxygenated Hb (double peak) and de-oxygenated Hb (single peak).
<br clear=right>
</p>
</td></table>

<p>There is a significant shift of the absorption peak in the 400-450 nm spectral range. However, we will focus on measuring the changes between 500-600 nm since the measurements are easier to perform and more reliable in this range. The absorption spectrum of oxygenated Hb exhibits two peaks in the 500-600 nm spectral range, while the spectrum of de-oxygenated Hb exhibits only a single peak. It is the change between these two states that you will quantify, and to do this we will focus on measuring diffusely reflected light from your palm and the inside of your forearm. These diffuse reflectance measurements will allow us to calculate the absorption spectra of Hb, correct for the effects of melanin from different skin types, and monitor the oxygen saturation state of Hb as we simulate a state of reactive hyperaemia, which is a brief increase in blood flow following a period of ischemia, or arterial occlusion.</p>
<br style="clear: both" />
<table width=160 align=left ><td>
<p align=justify>[[File:Specular_reflection.jpg|140px|border|center]]
<b>Figure 4</b> - Specular reflection from a surface.
<br clear=right>
</p>
<p align=justify>[[File:Diffuse_reflection.jpg|140px|border|center]]
<b>Figure 5</b> - Diffuse reflection from a multi-layer structure.
<br clear=right>
</p>
</td></table>

<p>To understand how we can make measurements of absorption by analysing diffusely reflected light, we should first define what is meant by the term reflection. In general light reflection can be defined in two ways; specular reflection, and diffuse reflection. Specular reflection refers to light that has been directly reflected from an interface, and is directional. A highly polished metal surface, such as a mirror, is an example of a specular reflector. This type of reflector will follow the law of reflection first described by Descartes, namely that the angle of incidence equals the angle of reflection (Figure 4). Another property of a specular reflector is that it will retain image information, which is why you can see your reflection in a mirror.</p>
<p>In diffuse reflection, the light can be thought of as penetrating a small distance into the reflector and scattering multiple times before exiting (Figure 5). This type of reflectance is non-directional, and does not produce any image since all image information in the wavefront is lost due to multiple scatterings. An example of a diffuse reflector would be a piece of white marble. No matter how much you polish the marble, most of the light striking its surface is diffusely reflected, which is why marble makes for a very poor mirror. A perfect diffuse reflector will reflect light uniformly into the 2π steradian space above it, while a perfect specular reflector will reflect light at an angle defined by the angle of incidence. In general most objects will reflect light both specularly and diffusely.</p>
<br style="clear: both" />

<p>Human skin can also be thought of an object which is diffusely reflective, like the marble, only that it also contains absorbers, namely Hb and melanin. Skin is a heterogeneous, multi-layered structure consisting of three basic layers, each containing numerous sub-layers. The basic layers of skin are the epidermis, which is the outermost layer and provides protection, the dermis, which serves as the location for hair follicles, sweat glands, etc, and the hypodermis, which consists of connective tissue to secure the skin to bones and muscle, as well as blood vessels to deliver oxygen and nutrients to the skin (Figures 6 & 7). Note that it is not important for you to memorize the various layers that make up the skin, but it is important to note where the chromophores we will be measuring reside and originate from. </p>
<br style="clear: both" />
<table width=360 align=left><td>
<p align=justify>[[File:Human_skin.jpg|320px|border|center]]
<b>Figure 6</b> - Cross section of human skin showing the major layers and components.
<br clear=right>
</p>
</td></table>
<table width=360 align=left><td>
<p align=justify>[[File:Skin_cross_section.jpg|320px|border|center]]
<b>Figure 7</b> - 3D representation of the skins layers and components.
<br clear=right>
</p>
</td></table>
<br style="clear: both" />

<p>Most of the diffuse reflection you will measure originates from the epidermal layer, which contains no blood vessels or capillaries. The blood diffuses through the dermal layer and into the epidermis, essentially meaning that there is a homogeneous distribution of Hb in the epidermal layer. For the purposes of this lab, you can consider the epidermal layer to be a perfect diffuse reflector, with a uniform distribution of melanin and Hb ‘absorbers’ present in a given volume. Since the diffusely reflected light is interacting with melanin and Hb as it is scattered within the epidermal layer, there is information within this light regarding the absorption properties of Hb and melanin.</p>

<h1>Methods </h1>
<p>To measure the diffuse reflectance spectra, we will make use of a ‘white’ LED which will be used as an illumination source. The diffuse reflectance will be collected with a fiber optic cable coupled to a computer controlled spectrometer. To simulate reactive hyperaemia, a compression cuff from a blood pressure monitor will be used to temporarily restrict blood flow to the hand. The Hb oxygen concentration will slowly drop following vascular occlusion, and immediately following re-perfusion, you will measure a significant jump in Hb oxygenation, then a steady return to normal physiological levels.</p>
<p>Before getting to the measurements, you should first familiarize yourself with the concept of a spectrometer and how the software interface should be utilized to obtain spectra with a good signal-to-noise ratio. A general schematic for the spectrometer used in this lab is shown in Figure 8.
<table width=360 align=left><td>
<p align=justify>[[File:Spectrometer.jpg|320px|border|center]]
<b>Figure 8</b> - Schematic of the light path in the Ocean Optics spectrometer used for this lab.
<br clear=right>
</p>
</td></table>
A spectrometer essentially consists of some light collection optics coupled to an optical fiber (not shown), the light collected by the fiber is passed through a lens and slit assembly mounted inside the spectrometer (2 & 3) before striking a collimating mirror (4). This mirror produces a collimated beam of light, which is then reflected off of a diffraction grating (5) and is focused by a second mirror (6) and onto a linear CCD array (7). A CCD (Charge-Coupled Device) is a light-sensitive detector which produces a voltage proportional to the light striking the active area, or pixels. The diffraction grating will reflect light of different colour at slightly different angles, and thus red light is focused towards the side of the CCD indicated by (8) and blue light is focused towards (9). Reading out the voltage levels on the pixels across the CCD therefore allows us to measure the spectrum of light collected by the fiber. When light is spread across the CCD in this fashion, the wavelength range striking each pixel on the CCD is very small, typically less than 0.25 nm per pixel, which allows for the visualization of very fine spectral features.</p>
<p>Software operation of the spectrometer will be quite basic for our purposes, the software is pre-installed on the PC you will be using for data collection, and can be found under the windows start menu at: Start → Ocean Optics → Spectra Suite. After initialization, the software will be running and ready to collect data. Ensure that you have connected the fiber optic cable between the spectrometer and light delivery/collection housing. The main parameters we will have to change are the exposure, averaging and boxcar settings. Typically it is good to use 4x averaging and 2-4x boxcar averaging. The exposure setting will vary based on the individual, but should be in the range of 200-1000 ms. The ideal gain setting will have the most intense pixel values at ~60,000 levels of grey (see Figure 9).</p>
<table width=560 align=center><td>
<p align=justify>[[File:Software_interface.jpg|520px|border|center]]
<b>Figure 9</b> - Spectra Suite software interface.
<br clear=right>
</p>
</td></table>
<p> Before we are ready to collect data we must first acquire several calibration data sets using the spectrometer to account for the natural spectral shape of the light source and optical system being used (this is the spectrum shown in Figure 10). This will be accomplished by measuring the diffuse reflectance of a ‘white’ reference target, which should have been supplied to you at the start of the lab. This target will allow you to measure the natural spectrum of the LED being used, and this data will later be used to normalize the diffuse reflectance skin spectra and remove any artefacts from the light source and fiber collection system.</p>
<table width=380 align=left><td>
<p align=justify>[[File:Save_spectrum.jpg|320px|border|center]]
<b>Figure 10</b> - Save Spectrum setup.
<br clear=right>
</p>
</td></table>
<p>collection system.
Once you have the proper gain setting for the reference spectrum, open up the save spectrum window (File → Save → Save Spectrum, or hold down “ctrl + alt + s”). The dialogue box shown in Figure *** should appear, and you should fill in the following settings:
</p>
<ol>
<li>Under “Save Options” select to “Save every scan”</li>
<li>Check the “Stop after this many scans” box, and enter “1” in the box beside it.</li>
<li>Under “File Type” select “Tab Delimited”</li>
<li>Select your “Save To Directory”</li>
<li>Enter “Reference” for the “Base Filename”</li>
<li>Ensure “Padding Digits” is set to “5”</li>
<li>Hit “Accept”</li>
</ol>
<br style="clear: both" />
<p>You should now have a single text file in your save to directory with the reference spectrum data. Now, remove the white reference target and place it back in its container. We are ready to begin spectral measurements. For this you will need the blood pressure compression cuff. Pressure to the cuff is increased by pumping on the bladder, and a silver release valve in front of the bladder allows the pressure to be released. To start the measurements follow the steps below:</p>
<ol>
<li>Open the “Manage Spectrum Exports” dialogue by either going to File → Save → Pause/Resume Export or pressing “ctrl + s”.</li>
<li>Highlight any processes and select “Terminate All”</li>
<li>Close the “Pause/Resume Export” dialogue</li>
<li>Open the “Save Spectrum” window</li>
<li>Under “Save Options” set it to save after every 2 scans</li>
<li>Check the option to “Pause until started by user”</li>
<li>Check the “Stop after this many scans” box, and enter “50” in the box beside it.</li>
<li>Under “File Type” select “Tab Delimited”</li>
<li>Select your “Save To Directory”</li>
<li>Enter “Back of Hand” for the “Base Filename”</li>
<li>Ensure “Padding Digits” is set to “5”</li>

<p>The dialogue box should look like this:</p>
<br style="clear: both" />
<table width=380 align=left><td>
<p align=justify>[[File:Acquisition_parameters.png|320px|border|center]]
<b>Figure 11</b> - Acquisition parameters for time-series measurements.
<br clear=right>
</p>
</td></table>
<br style="clear: both" />
<li>Hit “Accept”</li>
<li>Place the compression cuff just above your elbow, covering you bicep. Ensure that the pressure is fully released by opening the release valve for a few seconds then close it shut.</li>
<li>Place the reflectance scan head on the inside of your forearm (you should hold it in place while your lab partner operates the spectrometer software)</li>
<li>Optimize the exposure time so that the signal is maximized and no data channels are saturated</li>
<li>Ensure that you are using 4x averaging and 4x boxcar</li>
<li>Open the spectrum export dialogue by pressing “ctrl + s”</li>

<table width=380 align=left><td>
<p align=justify>[[File:Manage_spectrum.jpg|320px|border|center]]
<b>Figure 12</b> - Manage spectrum exports dialogue.
<br clear=right>
</p>
</td></table>
<br style="clear: both" />

<li>Highlight the process you just set up, which should show its status as “paused”. When you are ready to being, press “Resume All”. Data will now be collecting, and you should begin timing.</li>
<li>After 20-30 seconds, begin inflating the pressure cuff to capacity. This will restrict blood flow. The pressure should be maintained to at least 200 mmHg on the pressure dial.</li>
<li>After maintaining at least 200 mmHg for 90 seconds of constriction, release the pressure valve to allow blood to re-circulate.</li>
<li>Continue monitoring the directory you are writing spectral files into. When you reach file 49 you are done.</li>
<li>Press “Terminate All” to remove the completed data capture. If you forget to do this, you will not be able to set up another time series for the next set of measurements. If you find that this occurs, re-open the “Manage Spectrum Exports” dialogue will allow you to terminate the process and use the spectrometer again.</li>
</ol>
<p>After completing the above steps, repeat them while taking measurements of your lab partner’s inside forearm. After those measurements are completed, record spectra from the inside of your palm, then your partner’s palm. In total, you should have 4 complete data sets, but it may be a good idea to take 2 measurements from each site (8 in total between you and your partner). This way if you find some error during data processing, you have another set to analyze and won’t have to go back to the lab and run the experiment again.</p>
<p>Before leaving the lab please ensure that you have copied all of your data to a usb stick, or emailed it to yourself. You will also need to take with you a copy of the excel macro spreadsheet (.xlm file), which will be used to import all the spectra and calculate everything you need to quantify the change between oxy- and deoxy-haemoglobin. Finally, please ensure that the light source and spectrometer are properly shut down and stored away before you leave the lab.</p>

<h1>Data Analysis</h1>

<h2> Using the Excel Analysis Spreadsheet</h2>
<p> There is an MS Excel worksheet on the Desktop of the computer called "In-Vivo Spectroscopy Analysis (copy first).xlsm. This was made in Office 2007, but any version of Office should open the file and run the macros. You should set your macro security settings in Excel to run all macros. Copy this file and perform your data analysis using the copy.</p>
<ol style="list-style-type:lower-latin">
<li> Copy the Excel Spreadsheet and paste the copy in the directory you created for yourself to save you spectral data collected during the experiment</li>
<li> Open this Excel spreadsheet, you will notice there are six sheets of calculations, and 2 charts, but since you haven't imported the data yet, they are blank or show errors.</li>
<li> From the "Developer" tab at the top the window, select the "Macros" icon. </li>
<li> From the list which pops up, select the "ImportSpectralData" macros and click "Edit". </li>
<li> You will need to make the following three changes:
<ol>
<li> Change the entry for Data_Str1 to match the location where you collected your data (ie "Back of Hand000" or "Inside of Forearm000"</li>
<li> Change the entry for Spreadsheet to match the name you gave your copied version.</li>
<li> Change the "DirPath" entry to match the directory where your data is stored.</li>
</ol>
<li> Save the macro by clicking the "Save" icon, and return to Excel sheet by clicking the "Excel" icon</li>
<li> While looking at the Main Sheet, select the macro "ImportSpectralData" as before, and click "Run".</li>
<li> Data will be imported. It will take a couple of minutes for this process to complete. </li>
</ol>

<h2> Calculation Theory </h2>
<p> There are a number of steps that we must perform to normalize the data and convert the reflectance measurements into absorption measurements. These calculations are automatically set up and run in the excel macro for you, but you should understand the steps used and why they are used for answering questions during your experiment write-up. Below we will briefly run through the calculation theory. There are three main calculation steps taken to get the final data once it has been imported into excel. </p>

<ol>
<li>First the recorded spectra are normalized by the reference spectrum from the white reflectance target. This allows us to remove the spectral shape contribution of the light source, which can bias results if not accounted for.</li>
<li>Next these spectra must be converted from reflectance spectra into absorption spectra. This is accomplished via the equation shown below, which essentially states that the absorption spectrum of an object is logarithmically related to the inverse of the reflectance spectrum.</li>

<table width=200 align=center><td>
<p align=justify>[[File:iv_eqn1.png|200px|center]]
<br clear=right>
</p>
</td></table>

<li>The absorbance spectra are corrected for the effects of melanin by calculating the slope of the absorption spectrum in the region from 650 – 700 nm. This can be done because the absorption spectrum of melanin is essentially linear across the visible spectrum, and the absorption of haemoglobin is essentially flat in this spectral region, therefore any slope in this region is primarily due melanin concentration within the tissue. For each spectrum is calculated in this region, and the result of the linear fit is subtracted from the spectral data. This step effectively removes the melanin absorption spectrum from the data set.</li>
<li>Changes between oxy- and deoxy- states of haemoglobin are related to the single and double-peak spectra of these states. The change between these states can be quantified in a few ways. Here we will employ a very simple method which is based on calculating the slope over two regions in the spectra. This will give us a metric which will be used to quantify the change between oxy- and deoxy- haemoglobin (see figure below).</li>
</ol>
<table width=560 align=center><td>
<p align=justify>[[File:Sample_spectra.jpg|520px|border|center]]
<b>Figure 13</b> - Sample spectra of oxygenated and deoxygenated haemoglobin. Dashed lines indicate the slopes being calculated for quantification.
<br clear=right>
</p>
</td></table>
<p>The slope is calculated between 540-565 nm and 565-575 nm, and the difference between these two values gives a measure of the haemoglobin oxygen saturation. As can be seen in the figure above, when the blood is well oxygenated, the slope of the line segments are opposite, and at quite a large angle. In contrast, the slope of the two lines plotted on the de-oxy curve are almost the same value, and of the same sign. Therefore we would expect the value of H(t) to increase as the amount of oxygen is raised in the blood. The actual equation used is shown below, and as you can see it is quite simple mathematically, but it is important that you understand why it is valid to use such a simplified quantification model.</p>
<table width=200 align=center><td>
<p align=justify>[[File:iv_eqn2.png|250px|center]]
<br clear=right>
</p>
</td></table>

<p>This calculation is performed on each of the spectra you recorded, and the final step now is to generate a normalized haemoglobin oxygen concentration time series from this data. This is done by normalizing the ‘H(t)’ values in the above equation via:</p>

<table width=200 align=center><td>
<p align=justify>[[File:iv_eqn3.png|250px|center]]
<br clear=right>
</p>
</td></table>

<p>Plotting out H(t)norm as a function of time should then give a plot similar to the one shown below. From this plot you should be able to clearly see the steady decrease in oxygenation as the compression cuff was activated, and a dramatic spike in oxygenation following removal of the pressure and re-perfusion of the tissue with oxygenated blood.</p>

<table width=460 align=center><td>
<p align=justify>[[File:Haemoglobin_concentration.jpg‎|420px|border|center]]
<b>Figure 14</b> - - Haemoglobin concentration as a function of time during and after a period of vascular occlusion.
<br clear=right>
</p>
</td></table>

<h1>Questions/Write up</h1>
<p>The write-up for this lab does not have any specific formatting requirements. In general, you
should briefly describe the experiment and the experimental setup to give some background
information on what this lab intended to teach you (1-2 pages max). After this, you should
describe your data collection procedures as well as discuss your results in the context of the
questions asked of you below. The total report should not require more than 4-5 pages. If you
wish, you can simply answer the questions in the space provided below. Your write up should
answer or address the following questions and comments regarding the work you have done:</p>

<ol>
<li>Include the following plots with your final write up by taking a screenshot in excel and pasting them into your report. All of the values are calculated for you in the spreadsheet, but you must find the areas of max/min haemoglobin using the H(t)norm plot.
<ol style="list-style-type:lower-latin">
<li> H(t)<sub>norm</sub> vs time</li>
<li> I(t)<sub>max</sub>, I(t)<sub>min</sub>, I(t=10s), I(t=<sub>max</sub>)</li>
</ol>
</li>
<li>What is the main function of haemoglobin in the blood? Briefly describe the physical processes involved in haemoglobin’s biological functioning as well as what happens to the molecule in its various states of oxygenation.</li>
<li>How does the presence of melanin affect the data that you have collected and analyzed?</li>
<li>Briefly describe the difference between specular and diffuse reflection.</li>
<li>Briefly describe how a spectrometer works.</li>
<li>Why have we chosen to analyze absorption spectra by reflecting light off of a surface? What are the advantages of performing measurements such as these versus standard transmission spectroscopy?</li>
<li>Describe why the oxygen concentration increased so dramatically following the period of vascular occlusion. Is there anything you can say about how blood flows and oxygen is used up in the body on the basis of this data?</li>
<li>Can you suggest any improvements to this experimental setup that would make these measurements easier to perform in a clinical setting? What challenges would a clinical use of this technology face?</li>
<li>Why do we have to normalize our collected spectra with the spectrum reflected from the white reference target? What would happen to our measurements if we did not take this step?</li>
<li>Was there any difference in your results between the palm of your hand and the inside of your forearm? Please give an explanation for this difference, if you detected any.</li>
</ol>

Main Page/BPHS 4090/Optical Tweezers of Onions

2011-01-17T21:03:09Z

Sbilling: 91 revisions

<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: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]</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:onion1.jpg]]
[[ File:onion2.jpg]]
[[ File:onion3.jpg]]

<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:onion4.jpg]]
[[ File:onion5.jpg]]
[[ File:onion6.jpg]]
<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:onion7.jpg]]
[[ 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/Choloplast Translocation

2011-01-17T21:02:18Z

Sbilling: 55 revisions

<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>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>There is no 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 9 questions on the next page.</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>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>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 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>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>
<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>

<h1>References</h1>
<References/>

Main Page/BPHS 4090/microscopy II

2011-01-17T21:02:11Z

Sbilling: 28 revisions

<h1>Required Components</h1>
<ol>
<li>[[Media:Micrometer_Slide.JPG|Stage micrometer slide]]</li>
<li>[[Media:M2_Chroma_Slide_Kit.JPG|Chroma fluorescent slide kit]]</li>
<li>[[Media:M2_Bead_Slide.JPG|Fluorescent bead slide]]</li>
<li>[[Media:M2_Microchannel.JPG|Fluorescent microchannel slide]]</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 pipette 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>In this lab students will expand their use of optical microscopes to include fluorescence imaging,
which allows for highly specific labelling of proteins and complexes within the specimen.
Fluorescence is often complimented with Differential Interference Contrast imaging (DIC), and
this will also be explored during these experiments. Fluorescence really is a quantum
mechanical phenomenon, and the emphasis here will be on understanding what is physically
happening within samples that are fluorescently labelled. Understanding the principle of
fluorescence filtering and getting a feel for the light intensity levels being generated is critical for
fluorescence microscopy.</p>

<h1>Introduction</h1>
<p>Fluorescence microscopy is a very powerful tool allowing for the observation and quantification
of biological samples. You have seen previous labs how unstained samples contain very little
contrast, and through the addition of light absorbing dyes and stains or special optics, the
morphology and structure of samples can be readily observed. Fluorescence microscopy uses
the same concept of imparting artificial contrast to samples, but has several advantages over
absorption based contrast methods. In this contrast method, fluorescent molecules are
attached to specific structures within the specimen. Illuminating the specimen with a specific
wavelength of light causes these fluorescent molecules to convert some of this illumination into
a longer wavelength of light, which is then detected by a CCD or observed directly with the eye.</p>
<p>Fluorescence is a phenomenon first discovered by British scientist Sir George Stokes in 1852.
The process involves the absorption of an incident photon of light by a molecule, referred to as a
fluorophore, and subsequent emission of a second fluorescent photon a short time later (Figure
1). The energy transferred to the fluorophore after absorption causes the molecule to enter an
excited energy state that is short-lived, typically on the order of 10<sup>-9</sup> seconds (1/e value). During
this time, some of the incident energy is lost due to vibrational/rotational motion and heat,
among other things. When the molecule returns to the ground state, a fluorescence photon is
emitted at a longer wavelength (lower energy). This fluorescence can be detected with a colour
filter that passes the fluorescence emission and blocks the shorter wavelength excitation light.
The difference in wavelength between the incident and outgoing photons is referred to as the
Stoke’s Shift, and its phenomenon is the basis for all fluorescence imaging as it allows us to use a
colour filter set to separate the excitation and emission photons.</p>

<table width=500 align=center><td>
<p align=justify>[[File:Fl_fig1_states.png|480px|border|center]]
<b>Figure 1</b> –Schematic representation of the process of fluorescence. Light absorbed by a molecule causes it to move
into an electronic excited state. After losing some energy to heat/vibration/rotation, a fluorescent photon is
emitted and the system returns to the ground state.
<br clear=right>
</p>
</td></table>

<p>Fluorescent samples can be stained using two methods; dyes or antibody-based labels. Dyes
such as 4’-6’-Diamidino-2-phenylindole (DAPI) or Hoescht are commonly used in fluorescence
microscopy to stain nuclei. Both can be used on live or fixed cells, and are relatively
inexpensive. Upon incubation with a sample, DAPI binds with A-T rich repeats of chromosomes
in the DNA and the fluorescence emission efficiency increases versus its unbound state. This
enables visualization of cell nuclei when excited in the ultra-violet (UV) wavelength range and
produces blue fluorescent light. The chemical structure as well as absorption and emission
spectrum from DAPI are shown in Figure 2.</p>

<table width=400 align=center><td>
<p align=justify>[[File:Fl_fig2_spectrum.png|380px|border|center]]
<b>Figure 2</b> –Chemical structure and excitation/emission spectra of DAPI when bound to DNA. DAPI is excited by UV
light and produces blue fluorescent light. Its emission spectrum is quite broad in contrast to other fluorophores.
<br clear=right>
</p>
</td></table>

<p>To generate and detect the light emitted by fluorescent molecules some modifications to the
microscope light path are necessary (Figure 3). The excitation light is passed through an
excitation filter, which reduces the light to a specific wavelength, and is reflected off a dichroic
beamsplitter (DBS). A DBS is coated with a special surface designed to reflect light of one
wavelength range and transmit a different wavelength range. In the example shown in the
figure, the DBS will act as a mirror to blue light, and allow longer wavelength green light
generated at the sample to pass through it. The light then passes through one additional filter
(emission filter), which further separates any fluorescence from excitation light that may have
passed through the DBS.</p>

<table width=400 align=center><td>
<p align=justify>[[File:Fl_fig3_microscope.png|380px|border|center]]
<b>Figure 3</b> - Illumination path in an epi-fluorescence microscope. A filter cube contains an excitation filter, dichroic
beamsplitter, and emission filter and is responsible for separating the excitation light from the generated
fluorescence.
<br clear=right>
</p>
</td></table>

<p>The excitation and emission filters as well as DBS are contained within what is called a filter
cube, and these can be easily switched while working with the microscope to observe different
fluorophores. The microscope you will be working on has space for two separate filter cubes
operated on sliders. Other microscopes may use a wheel to translate the filters in and out of
the optical path. In highly automated microscopes these filter changes can be motorized and
controlled from the microscope software. The DBS directs light at the sample through epiillumination,
which means that the objective lens is used to deliver light to the sample and
collect the fluorescence emission. Recall that in transmitted light or brightfield imaging, a
condenser is used to illuminate the sample from the opposite side of the sample. Using epiillumination
to deliver excitation light through the sample allows for more efficient transfer of
excitation light to the sample (always a critical concern in fluorescence) and reduces the amount
of excitation light re-collected by the objective lens.</p>

<p>A typical configuration for a filter cube optimized for excitation and detection of DAPI is shown
in Figure 4 along with the excitation/emission spectra of DAPI. The excitation filter (blue line)
transmits light of a specific wavelength range which is efficient at exciting bound DAPI (blue
solid area). The DBS (green line) is designed to transmit wavelengths above ~ 410 nm, and thus
it will reflect almost all of the excitation light at the sample. The emission filter (red) is used to
clean up the light that is collected by the objective and passed through the DBS, and it is
optimized for detection of the emission peak of DAPI (red solid area). The emission filters used
in your microscope are longpass filters, which are slightly different from the bandpass filter
shown below. A longpass filter has a transmission spectrum similar to the tail of the DBS spectra
and passes all wavelengths above a certain threshold. This allows for collection of more of the
fluorescence signal which typically increases image quality, but prevents multiple fluorophores
from being imaged simultaneously.</p>

<table width=500 align=center><td>
<p align=justify>[[File:Fl_fig4_filter.png|480px|border|center]]
<b>Figure 4</b> - Typical filter cube configuration for detection of DAPI in stained specimens. The bandpass emission filter
(red line) can also be swapped out for a longpass filter, similar in shape to the tail of the DBS spectrum (green line).
<br clear=right>
</p>
</td></table>

<p>It is possible to use a single filter cube to visualize multiple fluorophores simultaneously, and
while this is often done, it is generally better to acquire fluorescence images one channel at a
time. A filter configuration for observation of a common set of fluorophores (DAPI/FITC/Texas
Red) is shown below in Figure 5. Note that the absorption and emission maxima have been
normalized in the plots below. The relative strength of fluorescence from fluorophores will vary
from channel to channel, which will typically require adjustment of the excitation light intensity
or CCD exposure time for different channels.</p>

<table width=500 align=center><td>
<p align=justify>[[File:Fl_fig5_triple.png|480px|border|center]]
<b>Figure 4</b> - Triple filter configuration for detection of fluorophores simultaneously. Bleedthrough of DAPI
fluorescence into the green FITC detection channel often requires images to be acquired individually. FITC and
Texas Red emissions are separated enough to be viewed simultaneously.
<br clear=right>
</p>
</td></table>

<p>Shown in the figure are the excitation (dashed lines) and emission (solid lines) of DAPI (blue),
FITC (green) and Texas Red (red). To attempt viewing these three fluorophores simultaneously a
filter cube would have an excitation filter that passes light in the purple (365 – 395 nm), cyan
(475 – 495 nm) and yellow (550 – 570 nm) wavelength regions. These bands are selected to
match up with the absorption spectra of the three dyes to be excited. This excitation light
would then reflect off of a broadband beamsplitter which reflects only 10-20% of the excitation
light to the sample. This is done to maximize the transmission of fluorescence photons back
through the filter cube as 80-90% of them will pass through the beamsplitter to be detected. To
separate the fluorescence emissions from each other, the emission filter would only pass light in
the blue (430 – 460 nm), green (503 – 538 nm) and red (600 – 640 nm) ranges. These ranges are
selected to overlap with the fluorescence emissions, and avoid detection of any reflected
excitation light collected by the objective lens. To help you visualize the light path a bit better
refer to Figure 6 below.</p>

<table width=600 align=center><td>
<p align=justify>[[File:Fl_fig6_schematic.png|580px|border|center]]
<b>Figure 6</b>– Triple excitation and emission light path in a fluorescence microscope. The dichroic mirror in this instance is a broadband mirror with 10-20% reflection and 80-90% transmission to maximize fluorescence detection efficiency. The portions of the excitation and emission spectra from DAPI/FITC/Texas Red contained in the filter bandpasses are shown as dashed and solid lines, respectively.
<br clear=right>
</p>
</td></table>

<p>On the excitation side we can see that the DAPI excitation actually causes some excitation of the FITC fluorophore. This can cause ghost images to appear if filtering is not properly performed. On the emission filter side we can see that there is no FITC emission in the DAPI detection channel (blue region), and thus the excitation of FITC by the DAPI excitation light is not detected. When examining the FITC emission channel however you can see that there is a significant amount of DAPI fluorescence (blue solid line) that would be passed by the FITC emission filter (green region). This is often a reason why individual excitation and emission filters are used; this issue goes away if no UV excitation light is striking the sample while observing FITC. In contrast, the distance between FITC emission (green line) and Texas Red emission (red line) is large enough that there is a minimal amount of FITC emission passing into the Texas Red detection channel (red region). The take-home message here is that simultaneous imaging can be performed more reliably when the emission spectra are better separated.</p>

<h2>Antibody Labelling</h2>
<p>While stains such as DAPI will bind to DNA, it stains the whole DNA strand rather non-specifically. Antibody labelling is another way to get fluorescent molecules coupled to the sample, and can be used to identify specific extracellular and intracellular proteins as well as specific mutations or activation regions within DNA. Different cell types express different antigens on their cell surfaces, and antibodies targeted against these antigens allow these cells to be distinguished from each other. While fluorescent molecules such as DAPI stain DNA fragments and yield morphological information on the specimen, the staining is more passive in that all DNA and some RNA is stained. Antibody labelling can be specific to sub-populations of cells or even highly specific regions of DNA. This imparts the ability to visualize the sub-cellular distribution of a wide range of biomolecules and investigate their functions at a molecular level.</p>

<table width=500 align=center><td>
<p align=justify>[[File:Fl_fig7_labelling.png|480px|border|center]]
<b>Figure 7</b>– Representation of a 2-step antibody labelling process. A primary antibody is first incubated with the
sample and labels the antigen of interest (left). In a second step, a fluorescent antibody recognizing the primary
antibody labels the cell (middle). After washing any unbound labels, fluorescence from the bound fluorescent
antibody can be detected and quantified (right).
<br clear=right>
</p>
</td></table>

<p>Figure 7 demonstrates the concept of fluorescent antibody labelling using a two-stage process
(secondary labelling). First, an antibody that is specific to a particular antigen expressed on a
cell is incubated with the specimen and binds to the target (left). The power of antibody
labelling lies in the fact that the shape of the antibody can only fit with its unique antigen, thus
after incubation and rinsing any unbound molecules are washed away. Next, a secondary
antibody linked to a fluorescent molecule is incubated with the sample and allowed to bind to
the primary antibody (middle). After rinsing and washing again the only fluorescent molecules
remaining in the sample are those attached to the target antigen through the antibody link.
Excitation/detection can then proceed by using the appropriate filter cube to visualize the
localization of the fluorescence signals.</p>

<p>Since multiple biomarkers can be localized in a single specimen through the use of secondary
antibodies that excite/emit at different wavelengths (taking cross-talk into consideration as
discussed above) fluorescence labelling is a really powerful tool for cell biology. This is because
in transmitted light imaging typically one or two labels can be applied to each slide, but with
fluorescence imaging the number of fluorescent probes (and thus antigens/proteins studied in
the same population of cells) can be significantly larger. A table of common fluorophores along
with their excitation/emission wavelengths are shown in the table below.</p>
<table width=400 align=center><td>
<p align=justify>[[File:Fl_fig7b_table.png|400px|border|center]]
<br clear=right>
</p>
</td></table>

<h2>Photobleaching</h2>
<p>In general, the fluorescence intensity emitted by a stained specimen is linearly related to the
light intensity used to excite the fluorescence. However, there are also probes available that
alter their fluorescence quantum efficiency or emission spectra based on the micro-environment
they are in. You will not be working with these types of probes in this lab as their use is too
advanced for the limited timeframe we have, but it is good to be aware they exist.</p>

<p>The process of fluorescence is inherently destructive to the probe molecules, and there is a limit
to how much light a given fluorophore can emit before it loses its ability to fluoresce. This
process is called photobleaching, and is generally the result when too much excitation power is
placed on the specimen or observation continues over extended periods of time. When this
occurs, the fluorescent molecules undergo a structural change which alters their ability to
produce more fluorescent photons. Instead, any energy absorbed by photo-destroyed
fluorophores is dissipated in a non-radiative fashion. A typical example of photobleaching is
shown in Figure 8.</p>

<table width=500 align=center><td>
<p align=justify>[[File:Fl_fig8_photobleaching.png|480px|border|center]]
<b>Figure 8</b>– Example of photobleaching as a function of exposure for a fibroblast cell. Over very long exposures
simulating prolonged observation, the nuclear fluorescence has gone away, and the green fluorescence has faded.
The red fluorescence is virtually unchanged.
<br clear=right>
</p>
</td></table>

<p>The specimen is a fibroblast cell, fluorescently labelled with three probe-antibody combinations; Hoechst 33258 (stains nuclei, excited by UV light, produces blue fluorescence), Alexa 488-phalloidin (excited by blue light, produces green fluorescence), and MitoTracker Red (stains mitochondria and cytoskeleton, excited by green light, produces red fluorescence). Images were acquired at 2-min intervals, and the level of fluorescence from the nuclear Hoechst stain can be seen to drop rapidly with exposure to the excitation light, while the signal from the Alexa 488 fluorophore drops only slightly, and the red fluorescence remains constant. Photobleaching complicates the routine use of fluorescence for quantitative studies, and care must be taken to minimize its effects. Often this involves minimizing the excitation intensity of light on the sample and detecting the fluorescence as efficiently as possible. Typically, fluorescently labelled specimens are imaged in a dark environment and are stored in a fridge or freezer protected from ambient light so their fluorescent properties are maintained. It is also very good practice to only illuminate the specimen when collecting data or observing the specimen through the oculars, then close the excitation shutter while saving data and making notes in your lab book (do not turn the lamp on and off however, as this can damage the bulb).</p>

<h2>Fluorescence Photon Efficiency</h2>
<p>A useful exercise to understand some of the finer points of fluorescence microscopy is to go through an estimation of the generation and detection of fluorescent light from a uniformly fluorescing sample. The excitation source is assumed, for this exercise, to be a standard 75 W xenon arc-discharge lamp having a mean luminous flux density of approximately 400 cd/mm<sup>2</sup>. When an excitation filter is used to pass blue light from 485 – 495 nm, about 2 mW of light will pass through to be directed at the sample by the DBS in the filter cube (assuming 75% transmission efficiency of the filter). Assume the DBS reflects 90% of this light towards the sample, and that gives a value of approximately 1.8 mW entering the rear aperture of the microscope objective as the excitation beam.</p>
<p>With a 100x/1.4 NA oil immersion objective the area of the specimen illuminated will be approximately 40 μm in diameter, giving an illumination area of 12 x 10<sup>-6</sup> cm<sup>2</sup>. The light flux on the specimen is then about 150 W/cm<sup>2</sup>, which corresponds to a flux density of 3.6 x 10<sup>20</sup> photons/cm<sup>2</sup>. This illumination intensity on the specimen is about 1000 times higher than that incident on the Earth's surface on a sunny day, yet with all this power only a fraction will generate fluorescence that is collected, passed by the optical filters and detected by your eye or the CCD camera.</p>
<p>The fluorescence emission that results from the light flux discussed above depends on the absorption and emission characteristics of the fluorophore, its concentration in the specimen, and the optical path length of the specimen. We are considering a uniformly fluorescing material, and in mathematical terms the fluorescence produced (F) is given by the equation:</p>
<table width=140 align=center><td>
<p align=justify>[[File:Fl_eqn1.png|140px|center]]
<br clear=right>
</p>
</td></table>
<p>Where σ is the molecular absorption cross-section, Q is the quantum yield, and I is the incident
light flux (estimated above to be 3.6 x 10<sup>20</sup> photons/cm<sup>2</sup>). Assuming that FITC is the fluorophore
being excited, σ = 3 x 10<sup>-16</sup> cm<sup>2</sup>/molecule, Q = 0.99, resulting in a value for F of 100,000
photons/sec/molecule. If the dye concentration is 1 μM/L and is uniformly distributed in a 40-
micrometer diameter disk with a thickness of 10 μm (volume = 12 pL), there will be
approximately 1.2 x 10<sup>-17</sup> moles (7.2 million molecules) in the optical path. If all of the molecules
were excited simultaneously with a bright pulse of light, the fluorescence emission rate will be:</p>
<table width=640 align=center><td>
<p align=justify>[[File:Fl_eqn2.png|640px|center]]
<br clear=right>
</p>
</td></table>

<p>The question of interest is how many of the emitted photons would be detected and for how
long could this emission rate continue? The efficiency of detection is mainly a function of the
optical collection efficiency (lens NA plus associated losses from filter cubes) and detector
quantum efficiency. A 1.4-numerical aperture objective with 100% transmission (an unrealistic
condition) has a collection efficiency of about 30%, which is actually quite high considering a
lens with a 90 degree collection volume (again, an unrealistic condition) would only collect 50%
of the generated fluorescence. The transmission efficiency of the DBS is typically 85% and the
emission filter will typically pass 80% of the in-band light. The overall collection efficiency is
therefore about 20%, or 1.4 x 10<sup>11</sup> photons/sec. If the CCD has a typical sensitivity, the quantum
efficiency is about 50% for the green FITC emission (at 525 nm), so the detected signal would be
7 x 10<sup>10</sup> photons/sec. Comparing this value to the fluorescence emission rate calculated above
shows that only 10% of the generated fluorescence actually contributes to the signal you are
seeing. Even with a perfect detector (again, an unrealistic assumption), only about 20% percent
of the fluorescence emission photons can be detected. If an objective with a lower NA is used
(the 100x used in this example has a very high NA), the detection efficiency again drops further.</p>

<p>Another important note to re-iterate here is the effect of photobleaching. For FITC in an
oxygenated saline solution each molecule can only emit about 36,000 photons before being
destroyed. In a deoxygenated environment, the rate of photodestruction diminishes about
tenfold, so approximately 360,000 photons/molecule can be produced. The entire volume of
fluorophores in our uniform sample contained 7.2 million molecules. Therefore, a minimum of
2.6 x 10<sup>11</sup> and a maximum of 2.6 x 10<sup>12</sup> fluorescence photons could be produced before
photobleaching starts to diminish the signal noticeably. Assuming the emission rate of 1 x10<sup>5</sup>
photons/sec/molecule calculated above, fluorescence could continue for only 0.3 to 3 seconds
before photodestruction begins. In the case where 10% of the photon flux is detected, a signal
of 7.2 x 10<sup>10</sup> photo-electrons/second would be obtained at the CCD detector.</p>

<p>The numbers above certainly convey that fluorescence imaging is inherently performed under
low-light imaging conditions, and care must be taken to reduce background and stray light from
being detected and washing out the fluorescence signal. Following the argument of this
example, if the CCD contains a 1000 x 1000 pixel array, this signal would be distributed over a
million pixels, or approximately 72,000 photo-electrons per sensor. For a CCD with 9 μm<sup>2</sup> pixels
the storage capacity is about 80,000 photo-electrons and the read-out noise is less than 10
electrons. The signal-to-noise ratio would then be largely determined by photon statistical
noise:</p>
<table width=340 align=center><td>
<p align=justify>[[File:Fl_eqn3.png|340px|center]]
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</p>
</td></table>

<p>In almost all cases, this high signal level could only continue for a very brief period of time
before photodestruction occurs, and it is ultimately impractical to run with excitation powers
high enough to photobleach this quickly, especially in live cell fluorescence imaging. The
compromise utilized by most microscopists to prolong the observation period is a 10-20x
reduction in the excitation light intensity so that only a fraction of the fluorophore molecules are
excited and subjected to photodestruction at a given time. Thus, the signal-to-noise ratio rarely
equals the theoretical maximum in this example, and typically ranges between 10 and 20 in
fluorescence microscopy.</p>

<p>One final note of caution with fluorescence microscopes has to do with proper care of the
excitation lamp. These light sources need 5-10 minutes to warm up and become stable before
they should be used. After turning on the power wait 5 minutes before pressing the second
push button on the power supply. Once you have turned the fluorescence lamp on, it should be
left on for the duration of the lab, and you should use the shutter to control when light reaches
the sample for imaging.</p>

<h2>DIC Imaging</h2>

<p>The image forming physics and optical light path in DIC microscopy are quite different from modes explored so far in these labs, and has the most in common with phase contrast imaging in that both techniques enhance visualization of the optical path of samples. Recall that in a phase contrast microscope optical path length changes create phase delays in light passing through transparent samples, and interference is used to translate these phase changes into visible intensity changes. In DIC imaging, optical path length gradients (in effect, the rate of change of the optical path length) are primarily responsible for contrast. Steep gradients in path length generate excellent contrast, and images display a pseudo three-dimensional relief shading, which is characteristic of the DIC technique. Regions having very shallow optical path slopes, such as those observed in extended, flat specimens, produce insignificant contrast and often appear in the image at the same intensity level as the background. DIC works on the principle of interferometry to gain information about the optical density of the sample. A relatively complex lighting scheme produces an image with the object appearing black to white on a grey background. This image is similar in appearance to that obtained by DPC microscopy but without the bright diffraction halo around areas where the index of refraction changes. DIC is often used in combination with fluorescence samples to add morphological context to fluorescence images (see Figure 9).</p>

<table width=500 align=center><td>
<p align=justify>[[File:Fl_fig9_dicfl.png|480px|border|center]]
<b>Figure 9</b>– DIC and fluorescence images of two cell types. Both cell types are visible in the DIC image. Only fluorescing cells are present in the fluorophore image.
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</p>
</td></table>

<p>DIC works by separating a polarized light wavefront into two orthogonally polarized mutually coherent parts which are spatially displaced (or sheared) at the sample plane by a Wollaston prism (Figure 10). A Wollaston prism consists of two calcite prisms cut at right angles and mounted so that their optic axes are perpendicular to one another. The direction in which the polarizations are separated is called the shear direction. The outgoing light beams diverge from the prism yielding two polarized rays, with the angle of divergence determined by the prisms' wedge angle and the wavelength of the light. The displacement between these two polarizations is exaggerated in the figure, typically it is much less than the resolution limit of the microscope on the sample (i.e., < 0.2 μm).</p>

<table width=300 align=center><td>
<p align=justify>[[File:Fl_fig10_optical_path.png|280px|border|center]]
<b>Figure 10</b> -Optical path through a DIC microscope. Light is split into two orthogonal polarizations by a Wollaston prism and focused onto the sample. A second Wollaston prism recombines the two beams so they can interfere and produce the DIC effect.
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</p>
</td></table>

<p>After passing through the sample and being collected by the objective, these two polarized wavefronts enter a second Wollaston prism which recombines them, causing them to interfere with one another. If each of these wavefronts were observed individually before recombining, they would each have the appearance of a brightfield image, i.e. only intensity variations due to absorption would be visible. These two images do not quite line up because of the illumination shear introduced by the first Wollaston prism. This means that instead of interference occurring between rays of light that passed through the same point in the specimen, interference occurs between rays of light that went through adjacent points which therefore have a slightly different optical path. This recombination of light causes "optical differentiation" of the optical path length.</p>

<p>The second Wollaston prism is also used to vary the phase bias offset between the two polarizations. This is accomplished by translating the prism, changing the relative path lengths through each of the calcite wedges. The effect of this in the final image is to go from a darkfield type image, where most of the light is destructively interfered, and results in a pseuso three dimensional, which is image typically associate with the DIC mode. In both of these images, the features you see are not due to absorption differences in the sample, nor are they topographically representative of how the sample actually ‘looks’ to your naked eye since much of the contrast is generated by optical path differences you cannot typically see. </p>

<p>Since DIC contrast depends in some part to preserving the polarization of the light waves interacting with the sample, the use of plastic slides or culture dishes can destroy the contrast of the sample since plastic destroys this polarization relationship between wavefronts. This is not an issue for phase contrast imaging. One disadvantage of phase contrast imaging in combination with fluorescence is that the phase ring etched into the objective actually blocks some of the light collected by the lens which reduces light collection efficiency when used for fluorescence collection. DIC objectives do not have this limitation, and for this reason it is typically this mode that is used in conjunction with fluorescence imaging where we have just seen how efficient photon detection is very important.</p>

<p>As with phase contrast imaging, there is a procedure you must follow for proper alignment of the DIC optics. Refer to Appendix 1 for this procedure.</p>

<h1>Methods</h1>
<p>A schematic of the microscope you will be using in this lab is shown in Figure 11. 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 DIC. When imaging in fluorescence it is good practice to turn off the room lights. This will help reduce detection of scattered light as well as dark-adapt your eyes to make the low level fluorescence signals easier to see visually. Find the location of the mercury light shutter and slider bars for moving the fluorescence filter cubes in and out of the optical path, and be sure to keep the shutter closed and only open it for brief periods during fluorescence imaging.</p>

<table width=400 align=center><td>
<p align=justify>[[File:Fl_fig11_nikon.png|380px|border|center]]
<b>Figure 11</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.
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</p>
</td></table>

<p> Alignment of the DIC transmission optics is covered in Appendix 1, and initially you will look at transparent cheek epithelial cells, which you worked with in the previous lab, and move on to fluorescently labelled samples. If you have time at the end of the lab and are interested you can switch back to the phase contrast condenser and come back to the samples to compare DIC and phase contrast. When working with DIC and fluorescence it is important to remember to remove the polarizer from above the objective as well as the Wollaston prism when switching from DIC to fluorescence, otherwise the fluorescent signal will be attenuated further and your image exposure times will be extremely long.</p>

<p>The first fluorescent samples you will look at are made up of polystyrene microbeads (0.02 – 4 μm in diameter) which have been infused with a mixture of fluorescent molecules. Samples such as these are often used as resolution and intensity standards for fluorescence measurement devices. One slide contains microchannels which contain mixtures of two fluorescent microbeads too small to be resolved optically (0.02 μm) in diameter, so the bead solution appears like a homogeneous fluorescing medium like the example discussed above. Using this slide you will see how different wavelengths of light excite each fluorophore, as well as using time-lapse imaging to observe photobleaching in action. The second microbead slide consists of a series of circular areas which contain different sized microbeads in mixtures and on their own. The diameter decreases from 4 μm beads down to 0.02 μm, in the first five areas and in area number six there is a mixture of all sizes. Using this slide as well as spatially calibrated images you will measure and verify the size of the microbeads. Since these fluorescent beads are designed for routine calibration and testing of fluorescence systems, they are less susceptible to photobleaching than stained slides. That being said, care should still be taken to store these slides in the dark and try to limit the amount of light exposure they receive so that they last longer. </p>

<p>The fluorescence tissue sections used are from a set of four fluorescently labelled slides designed to act as a reference standard for microscope performance. They are optimized for measuring specific performance parameters of instruments, including resolution, cube performance or spectral separation and camera sensitivity. All four slides are labelled with the same fluorophores: Hoechst (UV excitation, blue fluorescence), Alexa488 (blue excitation, green fluorescence), Cy3 (green excitation, orange/red fluorescence) and Cy5 (red excitation, far red/IR fluorescence). The microscope you are using does not have a filter for the Cy5 dye, however, red excited dyes like Cy5 are difficult to visualize in any case because the human eye’s sensitivity to light past 700 nm is very poor. CCD detectors, on the other hand, are sensitive well into the IR and with the proper filter set digital images of Cy5 are efficiently captured.</p>

<p>One major advantage of dyes in the red excitation regime is that they excite less tissue autofluorescence, which can be problematic for UV, blue and even green excitation wavelengths. Tissue autofluorescence is the term used to describe background autofluorescence from the tissue specimen itself. There are a number of compounds naturally present in specimens which fluoresce naturally in live as well as fixed tissues. Tissue processing and fixation is also known to increase tissue autofluorescence, and the end result is often the reduction (and sometimes inability) to detect the fluorescence from the antibody or dye label which has been added to the specimen. Tissue autofluorescence from live cells originate from the mitochondria and lysosomes, most specifically from coenzymes such as nicotinamide adenine dinucleotide (NADH) and flavins such as flavin adenine dinucleotide (FAD). The fluorescence produced by these compounds have very broad emission spectra that can overlap much of the visible spectrum, making it almost impossible to eliminate their influence in fluorescence images with traditional filtering methods. In live tissues, collagen and elastin contribute greatly to autofluorescence, and generally express more endogenous fluorescence than cells in culture. The set of four fluorescent slides all exhibit moderate amounts of autofluorescence, and are sections of rat liver, skin, gut (ileum) and pancreas.</p>

<p>The liver, after the skin, is the largest organ in the body and its job is to process a huge variety of nutrients for use by individual cells throughout the body. The primary cell type within the liver is the hepatocyte, which is responsible for protein synthesis and storage as well as detoxification. Nuclei are stained with Hoechst, peroxisomes are stained with Alexa488, sinusoidal endothelial cells are stained with Cy3, and actin filaments are stained with Cy5. The key labelling components of this slide are the peroxisomes. Peroxisomes are organelles present in almost all eukaryotic cells and contain enzymes which rid the cell of toxic peroxides. They are bound by a single membrane which separates their contents from the cytosol (the internal fluid of the cell) (Figure 12). Each peroxisome is labelled with a primary antibody to PMP 70 which is linked to Alexa488. This molecule is abundant in the membrane layer. The size of individual peroxisomes varies between about 200 – 400 nm in cross section, as such they can be used as a resolution standard for light microscopy. If you examine the slide using a high NA objective (oil 40x or oil 100x) it should be possible to see the peroxisomes as “donuts” within the cell, at lower NA they appear as round solid structures.</p>

<table width=300 align=center><td>
<p align=justify>[[File:Fl_fig12_peroxisome.png|280px|border|center]]
<b>Figure 12</b> -Basic structure of a peroxisome. With fluorescent membrane staining and high resolution optics they appear as donut shaped objects in fluorescence images. Autofluorescence and photobleaching can make them difficult to detect.
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</td></table>

<p>The skin section is stained with Hoechst for nuclei, Alexa488 for actin, Cy3 for elastin, and Cy5 for collagen. The skin is the largest organ in the body; it cushions the body from assault, is the primary surface for heat regulation, and is a primary sensory surface. In this section the most obvious cellular structure is the stratum corneum (surface of the skin) and constitutes the top ten or so cell layers visible in the section. Immediately beneath the epidermis is a thin though tightly defined layer of basement membrane, labelled with antibodies to collagen IV. This thin layer essentially separates the epidermis from the dermis. The dermis has a rich blood supply and constitutes the “elastic” part of the skin. This tissue was chosen as it represents one of the more difficult structures in the body to image; apart from the principal cellular structures defined above you should also see significant autofluorescence from the elastin. This is particularly evident when using excitation from the UV through blue excitation. In fact it is reasonable to use autofluorescence to collect a useful image of the elastin in skin or large blood vessels.</p>

<p>The gut section is stained with Hoechst for nuclei, Alexa488 for tubulin, Cy3 for laminin and Cy5 for actin. The gut is one of the primary absorptive surfaces in the body, and is made up of tiny folds of tissue called villi. Villi look like ‘fingers’ and line the interior wall of the gut, and their shape serves to increase the surface area of the interior wall. The apex of each epithelial cell in villi is covered with actin rich microvilli, about 400 nm long that should be easily defined with any high NA objective above 40X. Goblet cells which intersperse the epithelial cells can be seen to be full of mucus droplets, and appear as vacuoles under H&E staining (Figure 13). This is an excellent slide to use in conjunction with DIC as the droplets are not visible in the fluorescence images, and easily seen in DIC. Under the epithelium is the lamina propria, which is highly vascularised tissue lining the gastrointestinal tract.</p>

<table width=400 align=center><td>
<p align=justify>[[File:Fl_fig13_gut.png|380px|border|center]]
<b>Figure 13</b> – H&E stained villi in a gut section. The large surface area of the villi is comprised of epithelial cells which contain tiny microvilli on their interior surface (not visible in figure), as well as goblet cells, which are primarily responsible for mucus secretion and storage (arrow).
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</p>
</td></table>

<p>The pancreas is important for producing several hormones in the body, such as insulin, and enzymes which aid in the breakdown of carbohydrates, proteins and fats. The pancreas section is stained with Hoechst for nuclei, Alexa488 for actin, Cy3 for insulin, and Cy5 for laminin. Individual pancreatic islets are stained for insulin to highlight the beta cells. These structures, like the peroxisomes in the liver sample, should be visible as discrete structures at high NA. However, unlike the peroxisomes, the beta cell granules are stained throughout and do not appear hollow.</p>

<h1>Experiments</h1>
<p>Before beginning the experiments there is one useful tip in working with fluorescence images which you may find useful: false-colouring the image from the CCD displayed on the computer monitor. This feature changes the image displayed from a greyscale image into a colour palate selectable from within the Image -> Lookup Tables menu in ImageJ. This will match the fluorescence emission colours you see with your eye (to a certain extent) and make the digital image more pleasing to view in some cases. Use blue when looking at DAPI, green when looking at Alexa488 and red for Cy3.</p>

<table width=400 align=center><td>
<p align=justify>[[File:Fl_fig13b_imagej.png|380px|border|center]]
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</p>
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<p>Also, when working with fluorescent images acquired with different filter cubes it is sometimes desirable to combine individual images into a colour composite image. For example, if you acquire one image using the blue excitation (producing green fluorescence) and another image using green excitation (producing red fluorescence) it is desirable to produce a colour composite of these two images. For this to work properly care must be taken that the sample is not shifted on the microscope when switching filter cubes or the images will not overlay properly. To overlay images in ImageJ select ‘Merge Channels’ from Image -> Colour:</p>

<table width=400 align=center><td>
<p align=justify>[[File:Fl_fig13b_imagej2.png|380px|border|center]]
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</p>
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<p>Select the appropriate colour for the merge overlay (UV/DAPI excitation = blue, blue/Alexa488 excitation = green, green/Cy3 excitation = red). If there are only two fluorescence channels, select ‘none’ in the drop down menu. If you are overlaying a DIC image on top of the fluorescence image, select the DIC image as the greyscale overlay. Also, uncheck ‘Create Composite, and check ‘Keep Source Images’ to keep the individual image channels open. When you click ‘OK’, the images will be merged into a colour overlay which you can save.</p>

<table width=300 align=center><td>
<p align=justify>[[File:Fl_fig13b_imagej3.png|280px|border|center]]
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</p>
</td></table>

<p>Finally, before getting to the experiments, here are some tips and reminders for working with fluorescence samples and combining the use of fluorescence with DIC imaging:</p>
<ol>
<li> <ol style="list-style-type:lower-latin">
<li>Ensure the room lights are off when working with fluorescence images. For this lab it would be good to work with the lights off for the duration, and use a small desk lamp for writing notes and seeing what you are doing.</li>
<li>Be liberal with the shutter controlling whether excitation light is reaching the sample or not and try not to leave the sample exposed to excitation light for more than 5-10 seconds at a time unless you are moving around looking for areas to focus in on.</li>
<li>Be sure to block the transmitted light or turn the halogen lamp (not the fluorescence lamp) off while acquiring fluorescence images. You can also leave the lamp on and block the light by placing an object underneath the condenser to block light from reaching the sample. This may be easiest.</li>
<li>Before acquiring fluorescence images make sure to remove the top polarizer and DIC prism from the mount on top of the objectives.</li>
<li>Likewise, replace the polarizer when acquiring DIC images, also making sure that you remove the fluorescence filter cube from the light path.</li>
<li>Clean oil objectives and re-useable slides thoroughly when finished. Inspect the lens with one of the eyepieces to verify it is clean.</li>
</ol>
</li>
<li>Align DIC mode using the method in Appendix 1.
<ol style="list-style-type:lower-latin">
<li>Prepare another cheek sample and onion sample as you did in the previous lab and observe them in DIC using the 20x and 100x DIC objectives. Be careful not to transfer any oil onto the 20x objective.</li>
</li></ol>

<li>Fluorescence excitation and photobleaching of sub-resolution bead mixtures.
<ol style="list-style-type:lower-latin">
<li>This sample is quite bright compared to typical fluorescent specimens, and contains beads ideally excited by blue and green light. You will notice that the blue light will actually excite both dyes.</li>
<li>The microchannels contain mixtures of the two beads in the following ratios: 100:0, 75:25, 50:50, 25:75, 0:100.</li>
<li>Acquire images of each of five microchannels using blue and green excitation light and the 40x oil objective.
<ol style="list-style-type:lower-roman">
<li>Set the exposure of the CCD to the value required to see the brightest microchannel without saturation, and use this exposure for all images using this that filter. In other words, you will have two image sets containing five images/set. Each image set should use the same exposure setting.</li>
</ol>
</li>
<li>Calculate the relative bead mixture ratios from your data set (Note: you can do this part after acquiring the time-series and putting the slide back).
<ol style="list-style-type:lower-roman">
<li>Open the image sets from the blue and green excitation wavelengths in ImageJ.</li>
<li>Using the ‘Rectangular Sections’ tool in ImageJ, draw a box within the
fluorescent area of the data set.</li>
<li>Pressing ‘Control + M’ on the keyboard will measure the intensity and standard deviation of the area within the box and input the results into a text box for.</li>
<li>Using the measured intensity calculate the relative bead abundances in each of the microchannels by taking the intensity of a given microchannel and dividing it by the sum of the intensities from the green and blue excitation wavelengths (see below).
<table width=150 align=center><td>
<p align=justify>[[File:Fl_eqn4.png|150px|center]]
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</p>
</td></table></li>
<li>How do your values compare with the physical amount of beads present in each
sample?</li>
</ol>
</li>
<li>Using the appropriate excitation wavelength and exposure for the channels on the
edges (100:0 & 0:100 mixtures, blue and green excitation) set up a timed acquisition in
μManager (refer to the previous lab if you need a refresher on how to do this) with the
following parameters:
<ol style="list-style-type:lower-roman">
<li>Number of frames = 100</li>
<li>Interval = 30 seconds/frame</li>
</ol></li>
<li>Open the time series and create and image stack of the data set and save it (see
previous lab for instructions if you don’t recall how to do this).
<ol style="list-style-type:lower-roman">
<li>Using the ‘Rectangular Sections’ tool in ImageJ, draw a box within the
fluorescent area of the data set that contains a relatively uniform region of
fluorescence.</li>
<li>Pressing ‘Control + M’ on the keyboard will measure the intensity and standard
deviation of the area within the box and input the results into a text box for
further processing.</li>
<li>Switch to the next image in the stack by pressing the ‘>’ key, and measure the
intensity again by pressing ‘Control + M’.</li>
<li>Repeat until you have gone through all frames in the stack.</li>
<li>Save the results file as a text file, import it into a spreadsheet program (Excel is
installed on the PC if you wish to do it now, or you can take the files with you
and do it at home as well).</li>
<li>Create a plot of intensity versus time (remember, the frames are 30s apart) for
each of the data sets. What do you notice about the intensity as a function of
time? Does one of the data sets show a larger effect?</li>
</ol>
</ol>
<li>Liver slide
<ol style="list-style-type:lower-latin">
<li>Obtain the fluorescent liver slide.</li>
<li>Using the 20x objective, find some areas with bright staining and move to the 40x (oil)
and 100x objectives.</li>
<li>Try and observe the peroxisomes using the lower resolution objectives (10x/20x). Do
you see them as donuts or spots?</li>
<li>Using the 40x (oil) and 100x objectives acquire fluorescence images using all three filter
cubes, particularly focusing on getting good clean images of the peroxisomes, which
should look like small ring shaped objects with these objectives.</li>
<li>Save all images and clean the slide and put it away.</li>
<li>Peroxisome measurement and RGB image generation.
<ol style="list-style-type:lower-roman">
<li>Input the spatial calibrations you calculated previously for these objectives and
generate and RGB merged image using the method described above.</li>
<li>Measure the size of the peroxisome, which should be around 200-400 nm. Do
your measurements agree?</li>
</ol>
</li>
</ol>
<li>Gut slide
<ol style="list-style-type:lower-latin">
<li>Obtain the fluorescent gut slide.</li>
<li>Browse around the slide and locate some villi.</li>
<li>Acquire images using 10x and 20x objectives with all filter cubes as well as by sliding the
DIC polarizers and prisms in place and acquiring a DIC image (note: DIC is only possible
using the 20x and 100x objectives on this system).</li>
<li>Repeat with the 100x objectives.</li>
<li>It may be useful to take the DIC images first to make sure that you see some goblet cells
in the field of view on the camera.</li>
<li>Save all images and clean the slide and put it away.</li>
<li>Image manipulations in ImageJ.
<ol style="list-style-type:lower-roman">
<li>Produce a fluorescence/DIC overlay in ImageJ of the frames you acquired using
the method described above.</li>
<li>Using spatially calibrated images, measure the size of the microvilli on the
columnar epithelial cells. They should be approximately 400 μm in length, and
barely visible on the 100x objective.</li>
<li>Also measure the diameter of the mucus secretions in the goblet cells.</li>
<li>Both of the above measurements may be easier to perform on the original
greyscale data rather than the colour composite.</li>
</ol>
</li>
</ol>
<li><b>(OPTIONAL)</b> Fluorescence of microbeads
<ol style="list-style-type:lower-latin">
<li>Obtain the fluorescent microbead slide. There are six regions marked on the slide, the
first five contain beads of a single size, and the last contains a mixture of all bead sizes.</li>
<li>Unlike the previous slide, the beads in this slide are each stained with four fluorophores,
and thus will be visible using all filter cubes. These beads will be visible as discreet
objects with most of the objectives. Use the filter cube which produces the lowest CCD
exposure setting for this slide.</li>
<li>Acquire images of each of the six areas using the 10x, 20x, 40x (air & oil) & 100x
objectives.</li>
<li>Take DIC images of beads on the area containing the bead mixtures (20x & 100x).</li>
<li>After you are finished with the slide, clean it using the supplied cleaner and return it to
its light protected box.</li>
<li>Bead size calculation and comparison of NA collection efficiency
<ol style="list-style-type:lower-roman">
<li>Open the images in ImageJ and calibrate the spatial pixel size using the values
you calculated in the previous lab using the calibration slide. You can also recalculate
them using the calibration slide again if you don’t have the numbers
handy. Follow the procedure in the previous lab if you need to do this.</li>
<li>Zoom in to a visible bead in an image and using the ‘Elliptical Selection’ tool in
ImageJ draw a circle around the bead and press ‘Control + M’ to measure its
intensity.</li>
<li>Use the ‘Straight Line Selection’ tool in ImageJ to measure the diameter of the
bead.</li>
<li>Repeat this for five beads and calculate an average size and average intensity for
each of the five bead sizes. Do this for the 40x (air & oil) as well as 100x
objective images.</li>
<li>For the intensity values from the 40x air and 40x oil images, normalize the
intensity by the exposure setting used for each of the images. In other words, if
you used a 100 ms exposure for the 40x oil and 400 ms exposure for the 40x air:
<table width=280 align=center><td>
<p align=justify>[[File:Fl_eqn5.png|280px|center]]
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</p>
</td></table></li>
<li>What do you notice about the normalized intensity between the air and oil
objectives? Does this make sense?</li>
<li>What is the minimum objective required to resolve all of the beads? Does the
lens NA play a factor in this at all?</li>
</ol>
</ol>
<li><b>(OPTIONAL)</b> Skin slide
<li>Obtain the fluorescent skin slide.</li>
<li>Switch between filters and objectives while browsing around the slide. Of particular
note is to observe how bright the autofluorescence is and how it is present in virtually
all image channels. As discussed above most of this autofluorescence is due to elastin
within the tissue.</li>
<li>There is no need to acquire images of this sample, but if you have time and you wish to,
you can acquire and generate an RGB overlay using the objectives of your choosing.</li>
<li>Save all images and clean the slide and put it away.</li>
</ol>

<h1>Questions</h1>
<ol>
<li>What transmitted light mode is best used with fluorescence and why?</li>
<li>Describe why using a higher NA lens for fluorescence is advantageous.</li>
<li>Describe what photobleaching is, and steps that can be taken to minimize it.</li>
<li>Describe some advantages of fluorescence labelling.</li>
<li>In the bead slide, what objective performed the best at resolving all beads as well as giving
the strongest signal?</li>
<li>How did your exposure settings vary between the microchannel slide, microbead slide and
tissue section slides? Using these values, estimate the relative difference of fluorescence
emission rates from each of these samples.</li>
</ol>

<h1>Appendix 1 – DIC Optical Alignment</h1>
<p>First, start by making sure that you are using the DIC condenser on the microscope. If not, ask your lab instructor or get it from the wooden condenser box and replace the phase contrast condenser with it.</p>
<ol>
<li>1. The optical path of a DIC microscope and its main components are shown in the figure below. Alongside this figure are actual images of what the components look like when removed from the microscope.
<table width=450 align=center><td>
<p align=justify>[[File:Fl_a1_1.png|450px|center]]
<br clear=right>
</p>
</td></table>
</li>
<li>Using the figures above as a reference, locate the condenser polarizer and prism selection turret, as well as the objective polarizer/prism mount on the microscope. In general you can work with the condenser polarizer always in position. The objective polarizer and prism should be removed from the optical path when performing fluorescence imaging.</li>
<li>To start, remove the objective polarizer and prism (above the objective) from the optical path by sliding out the pin and prism block on the mount.</li>
<li>Switch to the 20x DIC objective and focus on an unstained slide (cheek cell, baboon eye, onion cell, etc) sample using regular transmitted light.</li>
<li>Set the microscope up for Kohler illumination (see the previous lab if you need a refresher).</li>
<li>At this point you want to align the two polarizers for maximum extinction (maximum darkness) when both are inserted into the optical path. This is also known as ‘crossing the polarisers’ since the maximum extinction condition arises when the transmission axes of the polarisers are orthogonal, or crossed.
<ol style="list-style-type:lower-latin">
<li>To do this, loosen the mounting pin for the prism assembly above the condenser which is located on the left side of the microscope, silver knob on the microscope frame just above where the prism assembly mounts into the scope. Using both hands slowly rotate the condenser until the minimum amount of light is visible.</li>
<li>Tighten the stage condenser back to lock it into position.</li>
</ol>
</li>
<li>Insert the objective prism into the optical path by pushing the slider to the left.</li>
<li>Now rotate the condenser prism into position for the objective you are using (i.e., ‘20’ for the 20x objective, and if using the 100x objective you would rotate the condenser to the ‘100’ setting).</li>
<li>Replace the eyepiece and observe your specimen again.</li>
<li>Turn the prism alignment knob all the way in (turning clockwise). At one extreme you will see the specimen become highly coloured, which is not the alignment you want for traditional DIC.</li>
<li>Rotating the knob in the opposite direction, you should get to a position (near the middle of travel) at which the specimen exhibits a high degree of contrast, and appears to be illuminated from an oblique angle. Moving a little bit further, you will notice that the direction of illumination apparently changes 180° (ie, from top-left to bottom-right).</li>
<li>The alignment you want to be at is either side of this central point, and the image should present itself with a pseudo-3D, or embossed, appearance. The specimen should now exhibit a high degree of contrast.</li>
<li>Double check the Kohler illumination.</li>
<li>Two final notes:
<ol style="list-style-type:lower-latin">
<li>The direction of the ‘shadowing’ observed is the sheer angle (the angle between the laterally sheared beam produced by the condenser prism). The distance between an adjacent light and dark spot is the sheer distance, which is typically set to match the resolution of the objective lens being used in green light. This is why you need to use a different condenser prism for the 20x and 100x objectives.</li>
<li>Finally, remember that the topographic look of the image is not directly related to sample thickness, but the rate of change of optical path length within the specimen.</li>
</ol>
</li>
</ol>

Main Page/BPHS 4090/microscopy I

2011-01-17T21:02:05Z

Sbilling: 48 revisions

<h1>Required Components</h1>
<ol>
<li>[[Media:M1_Diatom.JPG|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>One of the most critical points of working with immersion objectives is to be very careful
with how much oil you use and to 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 microscopes 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 we 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 bright image, trying to minimize
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 dobj), 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>