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Main Page/BPHS 4090/In-Vivo Spectrocopy

2011-10-31T01:15:12Z

HanaD:

<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.
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</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.
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</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.
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</p>
<p align=justify>[[File:Diffuse_reflection.jpg|140px|border|center]]
<b>Figure 5</b> - Diffuse reflection from a multi-layer structure.
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</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.
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</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.
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</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>Part A: In-Vivo Spectroscopy</h1>
<h2>Methods </h2>

<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.
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</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.
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</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.
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</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>
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<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.
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</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.
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</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>

<h2>Data Analysis</h2>

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

<h1>Part B: Cytochrome C</h1>

<p>Cytochrome is oxidized by potassium ferricyanide and reduced by sodium hydrosulfite.</p>
<p>We will be using the built-in Absorbance wizard of the SpectralSuite software to analyse the spectrum of cytochrome in the deoxygenated and oxygenated state.</p>
<p>The Absorbance wizard stores in its memory the spectrum of the sample under three different conditions. The first spectrum reading is of the phosphate buffered solution with the addition of either potassium ferricyanide or sodium hydrosulfite. This serves as the control sample. The second spectrum reading is of the sample without the illumination lighting—this therefore measures any ambient lighting that could interfere with the measurements. The final spectrum is of the buffered cytochrome solution with the addition of either potassium ferricyanide or sodium hydrosulfite. The Absorbance wizard stores in its memory the spectrum of the sample under these three different conditions. It then subtracts the first two sample readings from the third sample reading. In this way, the background spectrum is removed from the sample, leaving only the change in intensity of the different reflected wavelengths. A negative change in intensity indicates an absorption.</p>

<ol>
<li>Prepare the control sample, by pipetting 1 mL of phosphate buffered saline into a cuvette. It is important to always hold the cuvette by the top edge so that fingerprints do not distort the spectrometer reading of the sample.</li>
<li>Using the scoopula, add about 2 mg of potassium ferricyanide to the buffer within the cuvette. You can use the scale to measure the amount of potassium ferricyanide, however in practice it is only necessary to estimate the amount.</li>
<li>Mix the potassium ferricyanide into the buffer by slowly pipetting the solution in and out of the pipette a few times. Be careful to avoid producing air bubbles. </li>
<li>Place the cuvette containing the buffer and potassium ferricyanide solution into the cuvette holder of the spectrometer. Turn on the spectrometer illumination source.</li>
<li>Next, open the Ocean Optics SpectralSuite software.</li>
<li>Under “File” select “New” and then select “Absorbance Measurement”.</li>
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<p align=justify>[[File:iv_cyt_fig1.jpg‎|420px|border|center]]
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</p>
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<li>Select “Next” from the window that appears.</li>
<li>Adjust the “Integration Time” so that the entire spectrum appears within the Preview window.</li>
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<p align=justify>[[File:iv_cyt_fig2.jpg‎|420px|border|center]]
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</p>
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<li>Click “Next” and in the resulting window, click the light bulb. This stores the control spectrum in memory.</li>
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<p align=justify>[[File:iv_cyt_fig3.jpg‎|420px|border|center]]
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</p>
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<li><p>Click “Next”. Block the spectrometer illumination source from reaching the sample. It is important to NOT turn off the spectrometer illumination source, as turning it back on requires additional time to let it warm up. Click the light bulb in the window. This stores the background light in memory.</p>
<table width=460 align=center><td>
<p align=justify>[[File:iv_cyt_fig4.jpg‎|420px|border|center]]
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</p>
</td></table>

<p>Now prepare the experimental sample. This consists of the cytochrome within buffer with the addition of either potassium ferricyanide or sodium hydrosulfite. This step is performed last because a spectrum measurement of the experimental sample is wanted immediately after mixing the reagents together. (Over time, the effect of the reagents on the cytochrome wears off)</p></li>

<li>To prepare the experimental sample, pipet 1 mL of phosphate buffered saline into a cuvette. Using a new scoopula, add about 1 mg of cytochrome into cuvette. (Typically, this amount is approximated as the smallest reasonable amount of cytochrome that can be transferred using the scoopula). Mix using the same procedure as before (It is crucial to avoid air bubbles!).</li>
<li>Next, add about 2 mg of potassium ferricyanide and mix as before.</li>
<li>Immediately place the cuvette into the holder of the spectrometer. Select “Finish” on the window. This takes the final spectrum of the experimental sample and automatically subtracts the two background spectrums from before.</li>
<li>Repeat this entire procedure but in “Step 12” add 2 mg of sodium hydrosulfite instead of potassium ferricyanide.</li>

<table width=460 align=center><td>
<p align=justify>[[File:iv_cyt_fig5.jpg‎|420px|border|center]]
A typical absorption spectrum of cytochrome mixed with sodium hydrosulfite.
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</p>
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<h1>Questions/Write up</h1>

<p> The following questions should be answered in your notes.</p>

<ol>
<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>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>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>

<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>Briefly describe how a spectrometer works.</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>
</ol>

Main Page/BPHS 4090/Choloplast Translocation

2011-10-23T12:15:01Z

HanaD: Changed question section

<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>Answer the following questions in your lab notebook.</p>
<ol>
<li>An additional experiment was performed [8] by illuminating only half the cell at a time. The
results of this experiment are shown below:
<table width=700 align=center><td>
<p align=justify>[[File:Ct_fig11.png|700px|center]]
<b>Figure 11:</b> Half Cell Illuminations: This particular cell was illuminated under broad band-pass
irradiation with a photon flux of <b>1500μmol m<sup>-2</sup> s<sup>-1</sup>.</b> After an hour of irradiation, chloroplasts have
vacated the illuminated area.
<br clear=right>
</p>
</td></table>
<p>What can you conclude about these results? Is chloroplast translocation a protective
mechanism for the cell or rather an avoidance mechanism? Explain in a few sentences.</p></li>

<li>Imagine you had a source of UV light available and you wished to perform an additional
experiment by illuminating the cells with UV light as you did earlier with the blue and red bandpass
filters. Assume your UV light source emits at a peak wavelength of 250nm ± 10nm. Taking
into account the absorption spectra of chlorophyll shown in Figure 1 and that DNA absorbs light
at 260nm what would you expect to happen?</li>

<li>During this lab we examined the wavelength dependence of light on systrophe, however other
experiments [8] demonstrated there is also an intensity dependence as shown in the Figure
below:</li>

<table width=500 align=center><td>
<p align=center>[[File:Ct_fig13.png|500px|center]]
<b>Figure 13:</b> Intensity Dependence of Light on Incipient and Complete Systrophe.
<br clear=right>
</p>
</td></table>

<p>At a photon flux of around <b>2000μmol m<sup>-2</sup> s<sup>-1</sup></b> there is a dramatic increase in the number of
incipient systrophe. Remember incipient means the cells are in the process of undergoing
chloroplast translocation. The photon flux we receive from the sun is around <b>2000μmol m<sup>-2</sup> s<sup>-1</sup></b>.
What does this imply?</p></li>

<li>Photosynthetic organisms use wavelengths between 400nm and 800nm for photosynthesis.
From Figure 1, it can be seen that chlorophyll strongly absorbs blue and red light. Taking this
into account and your answers to questions 1 and 2 what role do you think systrophe plays in
photosynthesis? Are they related? Please discuss in a few sentences providing insights into
your thinking process.</li>
</ol>

<p>For the formal write-up for this experiment, briefly describe the purpose of the experiment, the
experimental set-up and the physics behind the biological process we’re observing, this will be your
introduction. You should type up your results into a table comparing the illumination trials using red
and blue filters. Provide a brief discussion interpreting your results. In addition you need to provide
answers to the following questions.</p>
<ol>
<li>The effect of systrophe is thought to be a defence mechanism for the cell by decreasing the
absorptive cross sectional area of chloroplasts within the cell. We can test this hypothesis by
using a simple mathematical model to represent the cell. Assume each cell of E. viridis in its
resting state is a sphere with a shell of chloroplasts along the periphery of the cell. In reality
chloroplasts are also distributed radially along the cytoplasmic strands within the cell (analogous
to the spokes of a bicycle wheel); however for simplicity we will only consider the outer shell of
chloroplasts. After the cell undergoes systrophe, the chloroplasts surround the nucleus in a
thicker shell. In the table below some data are provided of cellular dimensions for 3 arbitrary
cells before and after systrophe using time lapsed images <ref>Web-published: http://www.yorku.ca/planters/student_reports/viridis_systrophe.pdf (accessed
9 September 2010).</ref>.

<table width=500 align=center>
<tr><td><p align=center><b>Table 1:</b> Cellular Dimensions before and after Systrophe.</p></td></tr>

<tr><td><table width=500 align=center border>
<tr><td></td><td>Cell 1</td><td>Cell 2</td><td>Cell 3</td></tr>
<tr><td>Cell Diameter (μm)</td><td>122.5</td><td>108.5</td><td>130.5</td></tr>
<tr><td>Systrophe Diameter (μm)</td><td>80.1</td><td>64.7</td><td>103.8</td></tr>
<tr><td>Nucleus Diameter (μm)</td><td>41.8</td><td>45.8</td><td>61.3</td></tr>
<tr><td>Thickness of Chloroplast Shell
(Before Systrophe) (μm)</td><td>4.6</td><td>5.5</td><td>5.3</td></tr>
</table>

</td></tr></table>

<p>Use the data in Table 1 as well as the physical properties of the cells included in the
introductions and the Beer-Lambert Law (Equation 1.3) to fill in the table below:</p>
<table width=150 align=center><td>
<p align=justify>[[File:Ct_eqn1_3.png|150px|center]]
<br clear=right>
</p>
</td></table>
<p>Where I is the transmitted intensity of light passing through the cell, Io is the initial intensity your
light source, ε is the extinction coefficient of chlorophyll, C is the concentration of chlorophyll
and l is path length which the light passes through.</p>
<table width=700 align=center><td>
<p align=center><b>Table 2: </b> Absorptive Properties of Cells before and after Systrophe.
[[File:Ct_tab2.png|650px|center]]
<br clear=right>
</p>
</td></table>

<p>Based on your results in Table 2, does systrophe provide a protective mechanism for the cell by
decreasing the absorptive cross sectional area? Why would an increase in absorption
potentially be damaging for the cell? Consider both absorption and cross sectional area.</p>
</li>

<li>Open the spreadsheet titled <b>“Viridis_Systrophe_Spectrum.xls”</b>; this is an emission spectra of
the tungsten halogen bulb used for illuminating the cell. Calculate the total energy the cells
would receive from the whole spectrum of light emitted from this source by normalization. You
can use the spreadsheet to do your calculations and may wish to attach it to your lab report.</li>
<li>Below is the emission spectrum from the tungsten halogen bulb used for illuminations. From
the Planck Blackbody Distribution Law derive Wien’s Displacement Law which relates the colour
temperature of a blackbody to the peak wavelength it emits at. If we treat tungsten as a
blackbody, what is the color temperature associated with the peak wavelength observed in
Figure 12? Does this make sense considering the melting point of tungsten is about 3350K?
What correction do we need to take into account when deriving Wien’s Displacement Law from
Planck’s Blackbody Distribution Law to account for the unusual high melting point implied from
Figure 12?</li>

<table width=500 align=center><td>
<p align=center>[[File:Ct_fig12.png|500px|center]]
<b>Figure 12:</b> Tungsten Halogen Emission Spectrum.
<br clear=right>
</p>
</td></table>

<li>During systrophe chloroplasts are translated across the cell by molecular motors. It isn’t known
exactly which molecular motors are responsible for chloroplast translocation, however it should
be noted chloroplast translocation can be inhibited when cells of <i>E. viridis</i> are treated with 2,4 –
Dinitrophenol (DNP) [4]. The structure of DNP is shown in Figure 14.

<table width=400 align=center><td>
<p align=center>[[File:Ct_fig14.png|200px|center]]
<b>Figure 14:</b> Structure of 2,4 Dinitrophenol.
<br clear=right>
</p>
</td></table>
DNP acts as an uncoupling agent by disrupting the proton gradient the cell uses to create ATP.
Any energy the cell would have gained from ATP production is lost as heat. ATP provides the
energy for molecular motors to shuttle organelles across the cell. Based on the structure of
DNP, taking into account hydrophilicity and hydrophobicity and any partial charges on the
molecule explain how DNP inhibits chloroplast translocation. Does this provide any insight into
the energy mechanisms involved during systrophe?
</li>

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

Main Page/BPHS 4090

2011-09-14T12:19:00Z

HanaD:

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

<h2>Course Director</h2>
<p>Dr. Hana Dobrovolny</p>
<p>332 PSE</p>
<p>handob@yorku.ca</p>
<p></p>
<p></p>

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

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

<h1>Laboratory Manual</h1>
<ul>
<li> Sept. 19: [[Main Page/BPHS 4090/microscopy I|Transmitted Light Microscopy]] <b> </b> </li>
<li> Sept. 26: [[Main Page/BPHS 4090/microscopy II|Contrast Modes in Microscopy]] <b> </b> </li>
<li> Oct. 3: [[Main Page/BPHS 4090/Optical Tweezers of Onions|Optical Tweezers of Onion Cells]] <b></b> </li>
<li> Oct. 17: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 24: [[Main Page/BPHS 4090/Choloplast Translocation|Light Induced Chloroplast Translocation]] <b> </b> </li>
<li> Oct. 31: [[Main Page/BPHS 4090/In-Vivo Spectrocopy|In-Vivo Spectroscopy]]<b></b></li>
<li> Nov. 7: [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Nov. 14: [[Main Page/BPHS 4090/ElectroPhysiology of Chara revised|The Electrical Properties of ''Chara'']]<b> </b> </li>
<li> Nov. 21: [[Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy|Mapping a binding site using NMR spectroscopy]]<b> </b> </li>
</ul>

Main Page/BPHS 4090

2011-09-14T12:18:08Z

HanaD:

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

<h2>Course Director</h2>
<p>Dr. Hana Dobrovolny</p>
<p>332 PSE</p>
<p>handob@yorku.ca</p>
<p></p>
<p></p>

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

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

<h1>Laboratory Manual</h1>
<ul>
<li> Sept. 19 [[Main Page/BPHS 4090/microscopy I|Transmitted Light Microscopy]] <b> </b> </li>
<li> Sept. 26 [[Main Page/BPHS 4090/microscopy II|Contrast Modes in Microscopy]] <b> </b> </li>
<li> Oct. 3 [[Main Page/BPHS 4090/Optical Tweezers of Onions|Optical Tweezers of Onion Cells]] <b></b> </li>
<li> Oct. 17 [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Oct. 24 [[Main Page/BPHS 4090/Choloplast Translocation|Light Induced Chloroplast Translocation]] <b> </b> </li>
<li> Oct. 31 [[Main Page/BPHS 4090/In-Vivo Spectrocopy|In-Vivo Spectroscopy]]<b></b></li>
<li> Nov. 7 [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> Nov. 14 [[Main Page/BPHS 4090/ElectroPhysiology of Chara revised|The Electrical Properties of ''Chara'']]<b> </b> </li>
<li> Nov. 21 [[Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy|Mapping a binding site using NMR spectroscopy]]<b> </b> </li>
</ul>

Main Page/BPHS 4090/microscopy I

2011-09-10T02:33:31Z

HanaD:

<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>When working with immersion objectives be very careful with how much oil you use and clean the objective thoroughly when finished.</li>
<li>Also be careful not to get any oil on the non-immersion objectives. If this does happen,
remove the objective from the microscope and clean it with the lens cleaning solution.
Remove one of the microscope's eyepieces and use it to examine the lens and ensure it
is clean. Use the lens to look at a specimen to confirm it is clean. If it is still dirty, repeat
the cleaning procedure.</li>
<li>Move the stage down and use the light from the condenser as a reference for where to
place the drop on the specimen. You don’t need much immersion oil here, if you use
too much it will be a waste and make it more difficult to clean the slide when you are
finished.</li>
<li> Carefully finish rotating the 100x or 40x objective in place and re-focus on the sample.
At this point you can freely move around the slide, just try and keep the X-Y motion of
the sample to a slow and steady pace or else you will lose oil quickly and have to add
more. If you make slow, controlled motions the oil will not spread out as much.</li>
<li>After acquiring your images, lower the stage and remove the slide. Carefully clean the
slide and lens using a kimwipe and the supplied cleaning fluid.</li>
<li>When you are finished with the lab thoroughly clean any immersion objectives used as
well as slides that are part of the permanent lab (ie, micrometer, diatom slide, stained
slides). This is a critical step, as over time oil that is not cleaned can degrade image
quality and even render the very expensive objectives on the microscope useless. The
objectives will unscrew from the turret, and using one of the eyepieces you can examine
the lens after cleaning it to ensure there is no residual oil (ask your lab instructor to
show you how to do this if you are unsure).</li>
</ol>
<li><h3>Getting ready for acquisition and digital image capture.</h3>
<ol style="list-style-type:lower-latin">
<li>Turn the CCD on by pressing the power switch located on the top of the camera.</li>
<li>Launch the μManager software which will allow you to record data and adjust the
camera gain. Upon startup select the default configuration file in the splash screen that
starts up.
<table width=400 align=center><td>
<p align=justify>[[File:Fig7_micromanager.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>After startup, the main control screen for μManager should appear. From here you can
change the exposure settings on the CCD camera to optimize the signal by typing in an
exposure time (the units are milliseconds).
<table width=400 align=center><td>
<p align=justify>[[File:Fig8_config.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Rotate the eyepiece mounting block to the left to allow light to pass through to the CCD
camera.</li>
<li>Pressing the ‘live’ button in μManager should open a new window that shows a live
image of the light striking the CCD camera.</li>
<li>Pressing the ‘snap’ button will take a single snapshot which you can then save for later
processing.</li>
<li>Pressing the ‘Multi-D Acq’ button opens the multi dimensional acquisition dialogue from
which you can configure a time lapse acquisition (see below).</li>
</ol>
<li><h3>Setting up a timed acquisition in μManager</h3>
<ol style="list-style-type:lower-latin">
<li>For some experiments you will want to set up a timed acquisition so that you can make
a movie and dynamic measurements of samples. To do this open the multi-dimensional
acquisition dialogue and refer to the figure below to set relevant acquisition
parameters.
<ol style="list-style-type:lower-roman">
<li>Number = number of frames you want to acquire</li>
<li>Interval = time between frames (for setting up time-lapse)</li>
<li>Save Images = directory to save your images to (files will be automatically
numbered and timestamped)
<table width=400 align=center><td>
<p align=justify>[[File:Fig9_acquisition.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li></ol>
<li>It is a good idea to include parameters such as the CCD exposure time, microscope
objective, time delay and CCD bin settings in the file name or folder.</li>
<li>Be sure to change directories and file names between movies to keep the organization
of the files neat.</li>
<li>Pressing the ‘Acquire’ button will initiate the acquisition protocol using the parameters
you specified and the CCD gain currently being used.</li>
<li>Double check the folder after acquisition is complete to ensure all the files were
properly saved.</li>
</ol>
</ol>

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

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

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

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

<li><h3>(OPTIONAL) Using the diatom slide as well as the Pap smear slides compare the dry 40x objective with
the oil immersion 40x objective</h3>
<ol style="list-style-type:lower-latin">
<li>Focus on the sample using the 40x dry objective.</li>
<li>Switch to the ‘Live’ view on the CCD and adjust the exposure setting for the 40x dry
objective.</li>
<li>Rotate the dry objective out of the way and place a drop of oil on the slide and
rotate the 40x oil objective into place. Re-focus and observe the sample through
the eyepieces.</li>
<li>Do you see more detail with the one of the lenses? Explain why this is so.</li>
<li>Switch back to the ‘Live’ view on the CCD. What is the effect of NA on the CCD
exposure as well as image contrast?</li>
</ol>

</ol>

<h1>Questions & Tasks</h1>
<ol>
<li>Why do we use Kohler illumation? What is the major advantage of this illumination setup?</li>
<li>Briefly describe the difference between darkfield and brightfield and phase contrast
imaging.</li>
<li>How does the NA affect lateral (X-Y) resolution on images? Is there any effect along the Z
(focusing) axis?</li>
<li>Describe why oil immersion lenses produce sharper images as well as some of the things to
be cautious about when using immersion optics.</li>
<li>Calibrate the images you acquired and generate image stacks using the method described in
Appendix 3.</li>
<li>Calculate the cytoplasmic streaming rate in the onion cell using the method described in
Appendix 4 and compare with the value reported in the Stromgren-Allen & Brown paper.</li>
<li>You can create a movie of your cytoplasmic streaming data sets by saving your stack as an
AVI file within ImageJ (select ‘File’ -> ‘Save As’ -> ‘AVI’). Using a framerate of ~ 2 will show
the particles moving in real time. Also create movies running at 8 fps (4x real time) to see a
much more dynamic version of the streaming.</li>
</ol>

<h1>Appendix 1 - Background on Microscopy & Optics</h1>
<p>The most basic function of any microscope is to magnify an image and enhance the resolution
visible with the naked eye. In general, increasing magnification also increases the resolution
when the optical system being used has been designed and aligned properly. Before getting
into specifics, let’s review some fundamentals of how lenses and light form images.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig11_rays.png|500px|border|center]]
<b>Figure 1 -</b> Imaging properties of a single thin lens. The location and size of the image produced can be calculated
via the lens maker’s formula and the magnification ratio.
<br clear=right>
</p>
</td></table>

<p>The simplest way to produce a magnified image of an object is with a single lens, such as a
magnifying lens. Figure 1 shows a diagram that most of you should have seen at least once in
your university career, it is of a single ‘ideal’ thin lens and shows how light from the top of an
object is translated to an image plane by a lens with focal length ‘f’. Using the lens maker’s and
magnification formulas in the figure, the location and size of an object can be calculated if the
focal length and object size/position are known. When an object lies at a distance of 2f from the
lens, the image is formed at -2f and the magnification factor is -1 (ie, the image is inverted). As
the lens is moved closer to the object (decreasing d<sub>obj</sub>), the image distance and magnification
increase.</p>
<p>It is this image forming principle that is the basis for optical microscopes. The most basic form
of microscope, the compound microscope shown in Figure 2, contains two lenses. The objective
lens which collects light from the sample and the eyepiece lens, which further magnifies the
image generated by the objective and translates it to the back of the eye, or a detector such as a
CCD camera.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig12_compound.png|500px|border|center]]
<b>Figure 2 -</b> A simple compound microscope that can be used to produce a magnified image of a specimen. The
dotted line represents the apparent image size as seen by the observer or detector.
<br clear=right>
</p>
</td></table>
<p>In reality, each of these ‘lenses’ shown in Figures 1 & 2 are made up of a number of optical
components with differing shapes and glass compositions. A single lens is typically shown in the
figures to represent a multi-element lens, as diagrams can get very complex if this is not done.
The use of multiple lenses in a compound microscope allows a larger magnification factor to be
achieved, and also allows for corrections of image distortions (aberrations) which degrade image
quality when not properly corrected. A discussion of aberration correction and optical design is
beyond the scope of this lab and most undergraduate coursework, thus we will not go into this
in depth.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig13_na.png|400px|border|center]]
<b>Figure 3 -</b> Numerical Aperture (NA) is a measurement of the collection volume of a lens. In the figure, D represents
the lens diameter, f the focal length of the lens, n the index of refraction of the surrounding medium, and θ is the
maximum collection angle of the lens.
<br clear=right>
</p>
</td></table>
<p>Along with its magnification factor, the Numerical Aperture (NA) of an objective lens is very
important in reproducing sharp features in a specimen. The NA is a measure of the cone of light
collected from an objective lens; a higher NA indicates a lens that is more efficient at light
collection and produces a smaller focused spot size, which translates into better image contrast
and resolution. A diagram of the NA measure is shown in Figure 3. Light is focused to a point
which is one focal length from the lens centre. If the lens diameter is defined as D, then the
maximum collection angle of the lens for an on-axis point is then θ = tan-1(2f/D). The NA of a
lens is defined as NA = n x sin(θ), where n is the index of refraction of the medium between the
lens and the sample.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig14_focus.png|400px|border|center]]
<b>Figure 4 -</b> Focused spot size dependence on NA. As the NA is increased the collection volume increases and the
focused spot size gets smaller. This allows higher NA objectives to see finer details with better resolution and
sensitivity.
<br clear=right>
</p>
</td></table>
<p>As the NA is increased, the size of the focused spot decreases and the optical resolution
increases (Figure 4). Strictly speaking however, the distribution of the focused spot is not
actually a ‘spot’ but an Airy pattern, named after George Biddel Airy. This pattern arises due to
diffraction from the circular aperture of the lens, and the physical implication of this is that a
single point is not imaged into another single point, but into a distribution described by the Airy
pattern (Figure 5). The diameter of the Airy pattern (D<sub>Airy</sub>) can be calculated if the NA and
wavelength of light being used are known, and typically ½ of this value (Airy Radius) is defined as
the resolution limit of the system. Objects that are > 1 Airy unit apart will be clearly resolved as
two distinct points, while two points separated by < 1 Airy unit will blend together and be
indistinguishable from one another in the final image.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig15_airy.png|400px|border|center]]
<b>Figure 5 -</b> Images of two closely spaced objects as they approach the Airy resolution limit (A-C). Optical resolution
can be defined in terms of the width of the Airy pattern; two points separated by > 1 Airy radius will be resolved as
distinct objects, while objects closer than 1 Airy radius will blend together and appear as a single object (D).
<br clear=right>
</p>
</td></table>
<p>In practice, there are some fundamental limits to the magnitude of the NA possible. In the
equation for NA, sin(θ) is maximized when θ = 90°, but it is physically impossible to achieve this
collection angle in the real world. The largest NA possible for ‘dry’ objectives is around 0.95,
which corresponds to a collection angle of θ ≈ 72°. To further increase the NA beyond 0.95 a
‘wet’ or immersion objective is used. Increasing the index of refraction (n) of the region
between the lens and the sample also increases the NA, and typical immersion fluids are water
(n ≈ 1.33) and oil (n ≈ 1.55). While increasing the NA reduces the spot size, allowing smaller
features to be resolved, it also increases the light collection efficiency, thus improving the signalto-
noise ratio in images. Immersion optics are more difficult to work with, but they offer
unmatched resolution and image quality, as we will see during this lab.</p>

<h1>Appendix 2 - Phase Contrast Alignment</h1>
<p>This section gives an overview of the steps required to properly alight the phase plates in the
condenser and objective.</p>
<ol>
<li>Place a brightly stained specimen on the stage and rotate the 10x/Ph2 objective into the
optical pathway with the condenser on the BF setting. Follow the Kohler alignment
procedure from the main part of the lab.</li>
<li>Remove the sample and place a blank microscope slide on the stage.</li>
<li>Rotate the condenser turret into the Ph2 position.</li>
<li>Remove one of the microscope eyepieces and replace it with the phase telescope (if it is not
with the microscope, it will be in the wooden box that holds the DIC condenser).</li>
<li>While looking through the phase telescope, adjust its focus (slide it in and out of the tube)
until the phase plate in the objective is in sharp focus (it will look something like Figure 1a &
1c).</li>
<li>Locate the condenser annulus centering knobs and adjust the position of the annulus until it
is overlapped with the objective phase plate (Figure 1e). Note: do not attempt to adjust the
position of the condenser annulus with the main condenser centering knobs (the ones you
used for Kohler illumination). This effort will probably not achieve the intended condenser
annulus alignment and may compromise Kohler illumination.</li>
<li>The microscope should now be properly configured for phase contrast.</li>
<li>Take your prepared cheek cell sample and place it on the microscope.
<table width=500 align=center><td>
<p align=justify>[[File:Fig16_pc_align.png|400px|border|center]]
<br clear=right>
</p>
</td></table>
</ol>

<h1>Appendix 3 - Image stack generation and spatial calibration of images </h1>
<ol>
<li>Using images acquired as well as the stage micrometer images we will now generate an
image stack, which will allow you to quickly or slowly go through the data set frame by
frame. Also, the micrometer images will be used to spatially calibrate the data set.</li>
<li>Open the folder containing your data set and highlight all of the files you wish to open.</li>
<li>Drag all of the files onto the ImageJ program box (Important: to make sure the files import
in the correct numerical order make sure your mouse is on the first file when initiating the
drag and drop to ImageJ).</li>
<li>To turn the images into a stack click the ‘Image’ menu in ImageJ and select ‘Stacks’ then
‘Images to Stack’. This should combine all the separate image windows into a single
window. Moving the slider at the bottom left or right will play the sequence as a movie, or
you can go frame by frame.</li>
<li>Open the micrometer calibration image corresponding to the objective used to acquire the
stack that was just created.</li>
<li>Using the line tool in ImageJ (see below) draw a line across the marks on the slide. The lines
on the slide are separated by 10 microns, so measure from the leading edge of one black bar
to the leading edge of another bar across the field of view.
<table width=500 align=center><td>
<p align=justify>[[File:Fig17_imagej.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Open the ‘Analyze’ ImageJ menu and select ‘Set Scale’. The following window should pop
up, and already have the length of the bar you just drew (in pixels) filled in as the first entry.
Input the known calibration distance based on the number of dashes traversed by the line
and select the units to be microns. Also, check ‘Global’ to apply this calibration to all
opened images, including your stack.
<table width=200 align=center><td>
<p align=justify>[[File:Fig18_setscale.png|200px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Save and close your stack, then repeat with the rest of your images. Record the numbers for
each objective as you only have to measure the calibration once, and then can fill in the
numbers manually in the ‘Set Scale’ dialogue.</li>
<li>If you want to create a movie of your data stack select ‘File’ -> ‘Save As’ -> ‘AVI’. Selecting a
framerate of ~ 10 frames per second will be roughly 5x real time. These files can be played
in any computer media player.</li>
</ol>

<h1>Appendix 4 - Particle tracking and calculation of particle trajectories</h1>
<ol><li>Open an image stack from your onion cell data set taken in darkfield and ensure it is
calibrated properly.</li>
<li>Scan through the movie and try and identify a particle that stays in focus for at least 10
consecutive frames and can be clearly identified in each of them.</li>
<li>You can zoom in and out using the ‘+’ and ‘-‘ keys on the keyboard. You will find it easier to
track the particles if you zoom in a bit.</li>
<li>Select the ‘Segmented Lines’ tool from ImageJ by right clicking on the line tool icon as shown
below. This will allow you to left click on the particle in each frame and ‘draw’ its trajectory
on the stack. Double clicking will end drawing of the line.
<table width=400 align=center><td>
<p align=justify>[[File:Fig19_segmented.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Using the segmented line tool click on the centre of the particle you want to track. Using
the ‘<’ and ‘>’ keys on the keyboard you can scroll through the image stack frames. Track
your particle into the next frame and mark it again with a single left click, it should have
moved 10-12 microns. Repeat until the particle drifts out of view or becomes lost, ending
with a double click to finish the line draw procedure.
<table width=400 align=center><td>
<p align=justify>[[File:Fig20_stack.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>The number of boxes in the line segment indicates how many frames you tracked the
particle for. Count the number of frames you went through and call this number <charinsert>Δ</charinsert>f.</li>
<li>Now select ‘Analyze’ and then ‘Measure’ and a text box should pop up. The last entry in this
box should tell you the length of the line you just drew in microns. Call this <charinsert>Δ</charinsert>d.</li>
<li>To get the time taken, we need to confirm the time between frames from the metadata
which μManager exported while acquiring data. In the folder containing your data set there
should be a file called metadata.txt. Open this in wordpad and do a find for the highest
frame number acquired. Ie, if you set up the multi dimensional acquisition to take 100
frames, search for ‘100’. It should look like this.
<table width=400 align=center><td>
<p align=justify>[[File:Fig21_metadata.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Now, take the elapsed time (in ms) and divide it by the number of frames acquired to get
the time scale of your data set calibrated, call this value <charinsert>Δ</charinsert>t.</li>
<li>The average speed your particle is moving at is therefore:
<table width=100 align=center><td>
<p align=justify>[[File:Fig22_formula.png|100px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Repeat this calculation on multiple particles (~10) to get an average value from your data
set.</li>

Main Page/BPHS 4090/microscopy I

2011-09-10T02:19:44Z

HanaD:

<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>When working with immersion objectives be very careful with how much oil you use and clean the objective thoroughly when finished.</li>
<li>Also be careful not to get any oil on the non-immersion objectives. If this does happen,
remove the objective from the microscope and clean it with the lens cleaning solution.
Remove one of the microscope's eyepieces and use it to examine the lens and ensure it
is clean. Use the lens to look at a specimen to confirm it is clean. If it is still dirty, repeat
the cleaning procedure.</li>
<li>Move the stage down and use the light from the condenser as a reference for where to
place the drop on the specimen. You don’t need much immersion oil here, if you use
too much it will be a waste and make it more difficult to clean the slide when you are
finished.</li>
<li> Carefully finish rotating the 100x or 40x objective in place and re-focus on the sample.
At this point you can freely move around the slide, just try and keep the X-Y motion of
the sample to a slow and steady pace or else you will lose oil quickly and have to add
more. If you make slow, controlled motions the oil will not spread out as much.</li>
<li>After acquiring your images, lower the stage and remove the slide. Carefully clean the
slide and lens using a kimwipe and the supplied cleaning fluid.</li>
<li>When you are finished with the lab thoroughly clean any immersion objectives used as
well as slides that are part of the permanent lab (ie, micrometer, diatom slide, stained
slides). This is a critical step, as over time oil that is not cleaned can degrade image
quality and even render the very expensive objectives on the microscope useless. The
objectives will unscrew from the turret, and using one of the eyepieces you can examine
the lens after cleaning it to ensure there is no residual oil (ask your lab instructor to
show you how to do this if you are unsure).</li>
</ol>
<li><h3>Getting ready for acquisition and digital image capture.</h3>
<ol style="list-style-type:lower-latin">
<li>Turn the CCD on by pressing the power switch located on the top of the camera.</li>
<li>Launch the μManager software which will allow you to record data and adjust the
camera gain. Upon startup select the default configuration file in the splash screen that
starts up.
<table width=400 align=center><td>
<p align=justify>[[File:Fig7_micromanager.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>After startup, the main control screen for μManager should appear. From here you can
change the exposure settings on the CCD camera to optimize the signal by typing in an
exposure time (the units are milliseconds).
<table width=400 align=center><td>
<p align=justify>[[File:Fig8_config.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Rotate the eyepiece mounting block to the left to allow light to pass through to the CCD
camera.</li>
<li>Pressing the ‘live’ button in μManager should open a new window that shows a live
image of the light striking the CCD camera.</li>
<li>Pressing the ‘snap’ button will take a single snapshot which you can then save for later
processing.</li>
<li>Pressing the ‘Multi-D Acq’ button opens the multi dimensional acquisition dialogue from
which you can configure a time lapse acquisition (see below).</li>
</ol>
<li><h3>Setting up a timed acquisition in μManager</h3>
<ol style="list-style-type:lower-latin">
<li>For some experiments you will want to set up a timed acquisition so that you can make
a movie and dynamic measurements of samples. To do this open the multi-dimensional
acquisition dialogue and refer to the figure below to set relevant acquisition
parameters.
<ol style="list-style-type:lower-roman">
<li>Number = number of frames you want to acquire</li>
<li>Interval = time between frames (for setting up time-lapse)</li>
<li>Save Images = directory to save your images to (files will be automatically
numbered and timestamped)
<table width=400 align=center><td>
<p align=justify>[[File:Fig9_acquisition.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li></ol>
<li>It is a good idea to include parameters such as the CCD exposure time, microscope
objective, time delay and CCD bin settings in the file name or folder.</li>
<li>Be sure to change directories and file names between movies to keep the organization
of the files neat.</li>
<li>Pressing the ‘Acquire’ button will initiate the acquisition protocol using the parameters
you specified and the CCD gain currently being used.</li>
<li>Double check the folder after acquisition is complete to ensure all the files were
properly saved.</li>
</ol>
</ol>

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

<ol>
<li><h3>Calibration images of micrometer slide & setting CCD exposure</h3>
<ol style="list-style-type:lower-latin">
<li>Acquire and save images of the metric scale using the 10x, 20x, 40x (air and oil),
100x oil objectives.</li>
<li>Acquire the oil immersion images last, and ensure there is no oil on the slide when
using the dry objectives.</li>
<li>Please ensure the slide is cleaned thoroughly and placed back in its case after you
are finished with it.</li>
<li>Place the micrometer slide on the microscope and rotate the eyepiece turret
counter clockwise (to the left if facing the microscope) to allow the light to pass
through to the CCD camera.</li>
<li>Note the CCD camera has an additional 2.5x doubling lens in front of it. Thus a 10x
image to your eye is magnified up to 25x on the CCD. In other words; the field of
view you see with your eyes when using a 100x objective is what the camera sees
when using the 40x objective.</li>
<li>Press the ‘Live’ button and prepare to adjust the CCD exposure setting.</li>
<li>Adjust the CCD exposure setting so that you get 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 d<sub>obj</sub>), the image distance and magnification
increase.</p>
<p>It is this image forming principle that is the basis for optical microscopes. The most basic form
of microscope, the compound microscope shown in Figure 2, contains two lenses. The objective
lens which collects light from the sample and the eyepiece lens, which further magnifies the
image generated by the objective and translates it to the back of the eye, or a detector such as a
CCD camera.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig12_compound.png|500px|border|center]]
<b>Figure 2 -</b> A simple compound microscope that can be used to produce a magnified image of a specimen. The
dotted line represents the apparent image size as seen by the observer or detector.
<br clear=right>
</p>
</td></table>
<p>In reality, each of these ‘lenses’ shown in Figures 1 & 2 are made up of a number of optical
components with differing shapes and glass compositions. A single lens is typically shown in the
figures to represent a multi-element lens, as diagrams can get very complex if this is not done.
The use of multiple lenses in a compound microscope allows a larger magnification factor to be
achieved, and also allows for corrections of image distortions (aberrations) which degrade image
quality when not properly corrected. A discussion of aberration correction and optical design is
beyond the scope of this lab and most undergraduate coursework, thus we will not go into this
in depth.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig13_na.png|400px|border|center]]
<b>Figure 3 -</b> Numerical Aperture (NA) is a measurement of the collection volume of a lens. In the figure, D represents
the lens diameter, f the focal length of the lens, n the index of refraction of the surrounding medium, and θ is the
maximum collection angle of the lens.
<br clear=right>
</p>
</td></table>
<p>Along with its magnification factor, the Numerical Aperture (NA) of an objective lens is very
important in reproducing sharp features in a specimen. The NA is a measure of the cone of light
collected from an objective lens; a higher NA indicates a lens that is more efficient at light
collection and produces a smaller focused spot size, which translates into better image contrast
and resolution. A diagram of the NA measure is shown in Figure 3. Light is focused to a point
which is one focal length from the lens centre. If the lens diameter is defined as D, then the
maximum collection angle of the lens for an on-axis point is then θ = tan-1(2f/D). The NA of a
lens is defined as NA = n x sin(θ), where n is the index of refraction of the medium between the
lens and the sample.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig14_focus.png|400px|border|center]]
<b>Figure 4 -</b> Focused spot size dependence on NA. As the NA is increased the collection volume increases and the
focused spot size gets smaller. This allows higher NA objectives to see finer details with better resolution and
sensitivity.
<br clear=right>
</p>
</td></table>
<p>As the NA is increased, the size of the focused spot decreases and the optical resolution
increases (Figure 4). Strictly speaking however, the distribution of the focused spot is not
actually a ‘spot’ but an Airy pattern, named after George Biddel Airy. This pattern arises due to
diffraction from the circular aperture of the lens, and the physical implication of this is that a
single point is not imaged into another single point, but into a distribution described by the Airy
pattern (Figure 5). The diameter of the Airy pattern (D<sub>Airy</sub>) can be calculated if the NA and
wavelength of light being used are known, and typically ½ of this value (Airy Radius) is defined as
the resolution limit of the system. Objects that are > 1 Airy unit apart will be clearly resolved as
two distinct points, while two points separated by < 1 Airy unit will blend together and be
indistinguishable from one another in the final image.</p>
<table width=500 align=center><td>
<p align=justify>[[File:Fig15_airy.png|400px|border|center]]
<b>Figure 5 -</b> Images of two closely spaced objects as they approach the Airy resolution limit (A-C). Optical resolution
can be defined in terms of the width of the Airy pattern; two points separated by > 1 Airy radius will be resolved as
distinct objects, while objects closer than 1 Airy radius will blend together and appear as a single object (D).
<br clear=right>
</p>
</td></table>
<p>In practice, there are some fundamental limits to the magnitude of the NA possible. In the
equation for NA, sin(θ) is maximized when θ = 90°, but it is physically impossible to achieve this
collection angle in the real world. The largest NA possible for ‘dry’ objectives is around 0.95,
which corresponds to a collection angle of θ ≈ 72°. To further increase the NA beyond 0.95 a
‘wet’ or immersion objective is used. Increasing the index of refraction (n) of the region
between the lens and the sample also increases the NA, and typical immersion fluids are water
(n ≈ 1.33) and oil (n ≈ 1.55). While increasing the NA reduces the spot size, allowing smaller
features to be resolved, it also increases the light collection efficiency, thus improving the signalto-
noise ratio in images. Immersion optics are more difficult to work with, but they offer
unmatched resolution and image quality, as we will see during this lab.</p>

<h1>Appendix 2 - Phase Contrast Alignment</h1>
<p>This section gives an overview of the steps required to properly alight the phase plates in the
condenser and objective.</p>
<ol>
<li>Place a brightly stained specimen on the stage and rotate the 10x/Ph2 objective into the
optical pathway with the condenser on the BF setting. Follow the Kohler alignment
procedure from the main part of the lab.</li>
<li>Remove the sample and place a blank microscope slide on the stage.</li>
<li>Rotate the condenser turret into the Ph2 position.</li>
<li>Remove one of the microscope eyepieces and replace it with the phase telescope (if it is not
with the microscope, it will be in the wooden box that holds the DIC condenser).</li>
<li>While looking through the phase telescope, adjust its focus (slide it in and out of the tube)
until the phase plate in the objective is in sharp focus (it will look something like Figure 1a &
1c).</li>
<li>Locate the condenser annulus centering knobs and adjust the position of the annulus until it
is overlapped with the objective phase plate (Figure 1e). Note: do not attempt to adjust the
position of the condenser annulus with the main condenser centering knobs (the ones you
used for Kohler illumination). This effort will probably not achieve the intended condenser
annulus alignment and may compromise Kohler illumination.</li>
<li>The microscope should now be properly configured for phase contrast.</li>
<li>Take your prepared cheek cell sample and place it on the microscope.
<table width=500 align=center><td>
<p align=justify>[[File:Fig16_pc_align.png|400px|border|center]]
<br clear=right>
</p>
</td></table>
</ol>

<h1>Appendix 3 - Image stack generation and spatial calibration of images </h1>
<ol>
<li>Using images acquired as well as the stage micrometer images we will now generate an
image stack, which will allow you to quickly or slowly go through the data set frame by
frame. Also, the micrometer images will be used to spatially calibrate the data set.</li>
<li>Open the folder containing your data set and highlight all of the files you wish to open.</li>
<li>Drag all of the files onto the ImageJ program box (Important: to make sure the files import
in the correct numerical order make sure your mouse is on the first file when initiating the
drag and drop to ImageJ).</li>
<li>To turn the images into a stack click the ‘Image’ menu in ImageJ and select ‘Stacks’ then
‘Images to Stack’. This should combine all the separate image windows into a single
window. Moving the slider at the bottom left or right will play the sequence as a movie, or
you can go frame by frame.</li>
<li>Open the micrometer calibration image corresponding to the objective used to acquire the
stack that was just created.</li>
<li>Using the line tool in ImageJ (see below) draw a line across the marks on the slide. The lines
on the slide are separated by 10 microns, so measure from the leading edge of one black bar
to the leading edge of another bar across the field of view.
<table width=500 align=center><td>
<p align=justify>[[File:Fig17_imagej.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Open the ‘Analyze’ ImageJ menu and select ‘Set Scale’. The following window should pop
up, and already have the length of the bar you just drew (in pixels) filled in as the first entry.
Input the known calibration distance based on the number of dashes traversed by the line
and select the units to be microns. Also, check ‘Global’ to apply this calibration to all
opened images, including your stack.
<table width=200 align=center><td>
<p align=justify>[[File:Fig18_setscale.png|200px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Save and close your stack, then repeat with the rest of your images. Record the numbers for
each objective as you only have to measure the calibration once, and then can fill in the
numbers manually in the ‘Set Scale’ dialogue.</li>
<li>If you want to create a movie of your data stack select ‘File’ -> ‘Save As’ -> ‘AVI’. Selecting a
framerate of ~ 10 frames per second will be roughly 5x real time. These files can be played
in any computer media player.</li>
</ol>

<h1>Appendix 4 - Particle tracking and calculation of particle trajectories</h1>
<ol><li>Open an image stack from your onion cell data set taken in darkfield and ensure it is
calibrated properly.</li>
<li>Scan through the movie and try and identify a particle that stays in focus for at least 10
consecutive frames and can be clearly identified in each of them.</li>
<li>You can zoom in and out using the ‘+’ and ‘-‘ keys on the keyboard. You will find it easier to
track the particles if you zoom in a bit.</li>
<li>Select the ‘Segmented Lines’ tool from ImageJ by right clicking on the line tool icon as shown
below. This will allow you to left click on the particle in each frame and ‘draw’ its trajectory
on the stack. Double clicking will end drawing of the line.
<table width=400 align=center><td>
<p align=justify>[[File:Fig19_segmented.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Using the segmented line tool click on the centre of the particle you want to track. Using
the ‘<’ and ‘>’ keys on the keyboard you can scroll through the image stack frames. Track
your particle into the next frame and mark it again with a single left click, it should have
moved 10-12 microns. Repeat until the particle drifts out of view or becomes lost, ending
with a double click to finish the line draw procedure.
<table width=400 align=center><td>
<p align=justify>[[File:Fig20_stack.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>The number of boxes in the line segment indicates how many frames you tracked the
particle for. Count the number of frames you went through and call this number <charinsert>Δ</charinsert>f.</li>
<li>Now select ‘Analyze’ and then ‘Measure’ and a text box should pop up. The last entry in this
box should tell you the length of the line you just drew in microns. Call this <charinsert>Δ</charinsert>d.</li>
<li>To get the time taken, we need to confirm the time between frames from the metadata
which μManager exported while acquiring data. In the folder containing your data set there
should be a file called metadata.txt. Open this in wordpad and do a find for the highest
frame number acquired. Ie, if you set up the multi dimensional acquisition to take 100
frames, search for ‘100’. It should look like this.
<table width=400 align=center><td>
<p align=justify>[[File:Fig21_metadata.png|400px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Now, take the elapsed time (in ms) and divide it by the number of frames acquired to get
the time scale of your data set calibrated, call this value <charinsert>Δ</charinsert>t.</li>
<li>The average speed your particle is moving at is therefore:
<table width=100 align=center><td>
<p align=justify>[[File:Fig22_formula.png|100px|border|center]]
<br clear=right>
</p>
</td></table></li>
<li>Repeat this calculation on multiple particles (~10) to get an average value from your data
set.</li>

Main Page/BPHS 4090

2011-09-10T01:54:37Z

HanaD:

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

<h2>Course Director</h2>
<p>Dr. Hana Dobrovolny</p>
<p>332 PSE</p>
<p>handob@yorku.ca</p>
<p></p>
<p></p>

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

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

<h1>Laboratory Manual</h1>
<ul>
<li>[[Main Page/BPHS 4090/microscopy I|Transmitted Light Microscopy]] <b> </b> </li>
<li>[[Main Page/BPHS 4090/microscopy II|Contrast Modes in Microscopy]] <b> </b> </li>
<li>[[Main Page/BPHS 4090/Choloplast Translocation|Light Induced Chloroplast Translocation]] <b> </b> </li>
<li>[[Main Page/BPHS 4090/Optical Tweezers of Onions|Optical Tweezers of Onion Cells]] <b></b> </li>
<li> [[Main Page/BPHS 4090/In-Vivo Spectrocopy|In-Vivo Spectroscopy]]<b></b></li>
<li> [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> </b> </li>
<li> [[Main Page/BPHS 4090/ElectroPhysiology of Chara revised|The Electrical Properties of ''Chara'']]<b> </b> </li>
<li> [[Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy|Mapping a binding site using NMR spectroscopy]]<b> </b> </li>
</ul>

Main Page/BPHS 4090

2011-08-31T14:23:24Z

HanaD:

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

<h2>Course Director</h2>
<p>Dr. Hana Dobrovolny</p>
<p>handob@yorku.ca</p>
<p></p>
<p></p>

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

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

<h1>Laboratory Manual</h1>
<ul>
<li>[[Main Page/BPHS 4090/microscopy I|Transmitted Light Microscopy]] <b> Sep 20</b> </li>
<li>[[Main Page/BPHS 4090/microscopy II|Contrast Modes in Microscopy]] <b> Sep 27</b> </li>
<li>[[Main Page/BPHS 4090/Choloplast Translocation|Light Induced Chloroplast Translocation]] <b> Oct 4</b> </li>
<li>[[Main Page/BPHS 4090/Optical Tweezers of Onions|Optical Tweezers of Onion Cells]] <b> Oct 18</b> </li>
<li> [[Main Page/BPHS 4090/In-Vivo Spectrocopy|In-Vivo Spectroscopy]]<b> Nov 1</b></li>
<li> [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> Oct 25 / Nov 15</b> </li>
<li> [[Main Page/BPHS 4090/ElectroPhysiology of Chara revised|The Electrical Properties of ''Chara'']]<b> Nov 22 </b> </li>
<li> [[Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy|Mapping a binding site using NMR spectroscopy]]<b> Nov 29</b> </li>
</ul>

Main Page/BPHS 4090

2011-08-31T14:22:58Z

HanaD:

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

<h2>Course Director</h2>
<p>Dr. Hana Dobrovolny
handob@yorku.ca</p>
<p></p>
<p></p>

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

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

<h1>Laboratory Manual</h1>
<ul>
<li>[[Main Page/BPHS 4090/microscopy I|Transmitted Light Microscopy]] <b> Sep 20</b> </li>
<li>[[Main Page/BPHS 4090/microscopy II|Contrast Modes in Microscopy]] <b> Sep 27</b> </li>
<li>[[Main Page/BPHS 4090/Choloplast Translocation|Light Induced Chloroplast Translocation]] <b> Oct 4</b> </li>
<li>[[Main Page/BPHS 4090/Optical Tweezers of Onions|Optical Tweezers of Onion Cells]] <b> Oct 18</b> </li>
<li> [[Main Page/BPHS 4090/In-Vivo Spectrocopy|In-Vivo Spectroscopy]]<b> Nov 1</b></li>
<li> [[Main Page/BPHS 4090/Lysozyme Crystallization|Lysozyme Crystallization]]<b> Oct 25 / Nov 15</b> </li>
<li> [[Main Page/BPHS 4090/ElectroPhysiology of Chara revised|The Electrical Properties of ''Chara'']]<b> Nov 22 </b> </li>
<li> [[Main Page/BPHS 4090/Mapping a binding site using NMR spectroscopy|Mapping a binding site using NMR spectroscopy]]<b> Nov 29</b> </li>
</ul>