We thank the Light Microscopy Imaging Center at Indiana University. This work was supported by a grant from the National Institutes of Health (GM105755).

1. Preparation of necessary materials
<ol>
	<li>Prepare reagents for the growth and sporulation of yeast cells. 
	NOTE: If budding yeast strains are ade2- and trp1-, supplement all media in steps 1.1.1-1.1.3 with a final concentration of 0.01% adenine and 0.01% tryptophan from 1% stocks. If sterilizing media by autoclave, add these amino acids only after the reagents have been autoclaved and allowed to cool to room temperature.
	<ol>
		<li>For vegetative growth, prepare 2x synthetic complete + dextrose medium (2XSC) by dissolving 6.7 g of yeast nitrogen base without amino acids, 2 g of complete amino acid mix, and 20 g of dextrose in 500 mL of water. Sterilize the mixture by autoclaving for 20 min at 121 &#176;C or filtering through a 0.2 &#181;m filter.</li>
		<li>For the first step of sporulation, prepare 2x synthetic complete + acetate medium (2XSCA) by dissolving 6.7 g of yeast nitrogen base without amino acids, 2 g of complete amino acid mix, and 20 g of potassium acetate in 500 mL of water. Sterilize the mixture by autoclaving for 20 min at 121 &#176;C or filtering through a 0.2 &#181;m filter.</li>
		<li>For the final step of sporulation, prepare 1% potassium acetate (1% KAc) by dissolving 5 g of KAc in 500 mL of water. Sterilize the mixture by autoclaving for 20 min at 121 &#176;C or filtering through a 0.2 &#181;m filter.</li>
	</ol>
	</li>
	<li>Prepare drugs and reagents for microscopy, synchronization, and anchor away.
	<ol>
		<li>For adhering cells to the coverslip during time-lapse imaging, make 1 mg/mL of concanavalin A (ConA) in 1x PBS. Filter sterilize using a 0.2 &#181;m filter and store small (~10 &#956;L) aliquots at -20 &#176;C.</li>
		<li>To use the NDT80-in system, make 2 mL of 1 mM &#946;-estradiol dissolved in ethanol. Filter sterilize using a 0.2 &#181;m filter and store aliquots (~500 &#956;L) at -20 &#176;C.</li>
		<li>For the anchor away system, make 1 mg/mL of rapamycin dissolved in DMSO. Filter sterilize using a 0.2 &#181;m filter and store small (~10 &#956;L) aliquots at -20 &#176;C. 
		NOTE: Prepare small aliquots of rapamycin to avoid multiple freeze-thaw cycles of the drug.</li>
	</ol>
	</li>
	<li>Generate yeast strains containing the NDT80-in system (PGAL1,10-NDT80/PGAL1,10-NDT80; GAL4-ER/ GAL4-ER) and a target protein tagged with FRB in the anchor away genetic background (tor1-1/tor1-1; fpr1&#916;/ fpr1&#916;; Rpl13A-FKBP12/Rpl13A-FKBP12)4,8. Ctf19-FRB was used for this study. Additionally, one copy of histone protein Htb2 was tagged with mCherry to allow for monitoring of meiotic progression and chromatin segregation.</li>
	<li>Prepare a chamber for time-lapse imaging
	<ol>
		<li>Start preparing the chamber 6 h-24 h prior to imaging (Figure 2). Cut an 18 mm x 18 mm chamber from the pipette tip box insert using a heated scalpel or other sharp blade.</li>
		<li>Use a plastic pipette tip to spread a thin layer of silicone sealant around the bottom edges of the chamber that will adhere to the coverslip. Add enough sealant so that the edges of the chamber are completely covered (see Figure 2).</li>
		<li>Adhere the chamber to a 24 mm x 50 mm coverslip by gently placing the chamber, sealant-side-down, onto the coverslip. Make sure that there are no gaps in the sealant so that the chamber does not leak.</li>
		<li>Spread 8-10 &#181;L of ConA onto the coverslip. Dispense ConA onto the middle of the coverslip and use a pipette tip to spread the ConA into a thin layer such that it covers most of the coverslip that is surrounded by the chamber. 
		&#8203;NOTE: Plastic chambers can be reused indefinitely. After time-lapse imaging is completed, remove the chamber from the coverslip using a razor, clean the silicone sealant from the chamber, and keep the chambers submerged in 95% ethanol for future use.</li>
	</ol>
	</li>
</ol>
2. Sporulation of yeast cells
<ol>
	<li>Perform the following steps for starvation of yeast cells to induce the meiotic program. 
	NOTE: These steps are for the sporulation of the W303 strain of yeast. Other strains may require different protocols15. Sporulation efficiency is highly variable between lab strains, with W303 exhibiting a sporulation efficiency of ~60%16,17,18.
	<ol>
		<li>Take a single colony of the appropriate diploid yeast strain from a plate and inoculate 2 mL of 2XSC and let grow at 30 &#176;C on a roller drum for 12-24 h to saturation.</li>
		<li>Dilute the saturated culture into 2XSCA by adding 80 &#181;L of the culture from step 2.1.1 to 2 mL of 2XSCA. Let it grow at 30 &#176;C on a roller drum for 12-16 h. Do not leave in 2XSCA for longer than 16 h because cells can become sick or auto-fluorescent.</li>
		<li>Perform two washes by spinning down the culture at 800 x g at room temperature (25 &#176;C) for 1 min, discarding the liquid, and resuspending the pellet in 2 mL of sterile distilled water.</li>
		<li>After the second wash, remove the liquid and resuspend the pellet in 2 mL of 1% KAc and let it grow at 25 &#176;C on a roller drum for 8-12 h. Cells that have entered meiosis will arrest in pachytene of prophase I.</li>
	</ol>
	</li>
	<li>Prophase I release system
	<ol>
		<li>Add &#946;-estradiol directly to the sporulation culture to a final concentration of 1 &#956;M and vortex the culture tube quickly. The &#946;-estradiol will release cells from prophase I.</li>
	</ol>
	</li>
</ol>
3. Depletion of target protein from the nucleus using the anchor away technique
<ol>
	<li>Add rapamycin to the cells to deplete the FRB-tagged protein from the nucleus at a particular stage.
	<ol>
		<li>To deplete proteins from the nucleus at prophase I exit, add rapamycin to a final concentration of 1 &#956;g/mL to the sporulation culture tube at the same time as &#946;-estradiol addition.</li>
		<li>To deplete proteins from the nucleus at a particular stage of meiosis, monitor the cell cycle stage starting at 60 min after &#946;-estradiol addition. Every 20 min, pipette 5 &#181;L of culture onto a 24 mm x 40 mm coverslip and cover the cells with an 18 mm x 18 mm coverslip. Keep cultures spinning on the roller drum at 25 &#176;C between time points.</li>
		<li>Image the cells using the A594/mCherry filter and the 60x objective of a fluorescence microscope. Set the percent transmittance to 2% and the exposure time to 250 ms. The presence of one, two, or four DNA masses will indicate the meiotic stage at which the cells have progressed. Add rapamycin to a final concentration of 1 &#956;g/mL at the meiotic stage of interest.</li>
	</ol>
	</li>
	<li>Keep cells spinning on the roller drum at 25 &#176;C until they are ready for imaging. Nuclear depletion of a target protein occurs 30-45 min after rapamycin addition2,8.</li>
</ol>
4. Time-lapse fluorescence microscopy
<ol>
	<li>Make an agar pad that will be used to create a monolayer of cells for imaging (see step 4.2).
	<ol>
		<li>Cut off and discard the cap and bottom 1/3 of a 1.5 mL microfuge tube to create a cylinder. The cylinder will serve as a mold for the agar pad. Make two cylinders to have an extra in case the agar does not polymerize properly in the first one.</li>
		<li>Place the cut microfuge tube cylinder on a clean glass slide with the top of the tube upside down sitting on the slide.</li>
		<li>Make 6 mL of a 5% agar solution (use 1% KAc as solvent) in a 50 mL beaker and microwave it until the agar is fully dissolved. 
		NOTE: The agar will easily boil over in the microwave; watch the agar and start and stop the microwave when the agar starts to boil and swirl the beaker. Start and stop the microwave several times to fully dissolve the agar. The agar solution is made in excess of the amount needed to ensure that the agar dissolves fully.</li>
		<li>Cut off the tip of the pipet to make a larger opening and pipet ~500 &#181;L of melted agar into each microfuge tube cylinder. Let it sit at room temperature until the agar solidifies (~10-12 min).</li>
	</ol>
	</li>
	<li>Preparing yeast cells for imaging
	<ol>
		<li>Spin down 200 &#181;L of the sporulation culture from step 3.2 at 800 x g for 2 min in 1 mL microcentrifuge tubes. Remove and discard 180 &#181;L of the supernatant. Resuspend the pellet in the remaining supernatant by swirling and flicking the tube.</li>
		<li>Pipet 6 &#181;L of the concentrated cells onto the coverslip in the middle of the chamber made in step 1.4.</li>
		<li>Hold the cylinder with the agar pad made in step 4.1 and carefully slide it off the glass slide. Make sure the agar is completely flat at the bottom and using the bottom of a pipette tip, apply a slight amount of pressure to the microfuge mold such that the agar pad is pushed out slightly above the boundary of the tube.</li>
		<li>Invert the mold such that the agar pad is facing down toward the chamber.</li>
		<li>Using forceps, gently place the agar pad (still in the microfuge tube mold) on top of the cells. Use a pipette tip to gently slide the agar pad around the chamber 10-20 times to create a monolayer of cells on the coverslip.</li>
		<li>Leave the agar pad in the chamber for 12-15 min. This step will allow the cells to adhere to the ConA on the coverslip.</li>
		<li>Transfer 2 mL of sporulation culture from step 3.2 to two microcentrifuge tubes and spin at 15,700 x g for 2 min. Transfer the supernatant into clean microfuge tubes and spin again at 15,700 x g for 2 min. Transfer the supernatant to clean microcentrifuge tubes to be used in the next step. 
		NOTE: It is important to use the pre-conditioned KAc supernatant with the &#946;-estradiol and rapamycin to ensure efficient sporulation and continued depletion of the protein of interest.</li>
		<li>After the agar pad has sat in the chamber for 12-15 min, float and remove the agar pad before imaging. To do so, add 2 mL of the supernatant from step 4.2.7 dropwise to the chamber. Once the liquid has reached the top of the chamber, the agar pad will most likely float. 
		NOTE: If the agar pad does not float automatically, wait 1-2 min. If the agar pad still does not float, remove it gently with forceps. Ideally, the agar pad would float on its own because removing it prior to floating could lead to removal of the cells.</li>
		<li>After the agar pad has floated, remove it gently with forceps and discard it. Place a 24 mm x 50 mm coverslip on the top of the chamber to prevent evaporation during imaging.</li>
	</ol>
	</li>
	<li>Setting up a movie on the microscope 
	NOTE: The instructions below are for an inverted microscope fitted with a slide holder that accommodates the 24 mm x 50 mm coverslip (See Table of Materials for microscope, camera, and software details). A 60x oil-immersion objective is used for image acquisition. This protocol may need to be altered when using other microscopes or other slide holders. The exact steps for executing step 4.3.2 through step 4.3.16 will vary depending on the microscope and imaging software used. See section 4.4 for instructions for a different microscope.
	<ol>
		<li>Fit the coverslip inside the slide holder. Adhere the molding clay to the side of the coverslip to keep it secure in the slide holder.</li>
		<li>Open the image acquisition software. Use course and fine adjustment knobs to focus the cells using DIC or brightfield.</li>
		<li>On the main menu of the image acquisition software, click on File &#62; Acquire (Resolve 3D). Three windows will pop-up.</li>
		<li>In the window named Resolve 3D, click on the Erlenmeyer Flask icon. This will open a window titled Design/Run Experiment, which contains the controls for setting up an experiment to set up a time-lapse movie.</li>
		<li>Under the Design tab, navigate to the tab labeled Sectioning. Select the box next to Z Sectioning. Set the z stacks as follows: Optical section spacing = 1 &#181;m, Number of optical sections = 5, Sample thickness = 5.0 &#181;m.</li>
		<li>Under the Channels tab, click on the + icon such that one channel option appears. Select the appropriate channel. For this experiment, we select A594, which is used to image mCherry. Select the box next to Reference Image and set the Z position to the middle of the sample from the drop-down menu.</li>
		<li>Select a value from the dropdown menu adjacent to %T and Exp. to set percent transmittance and exposure time, respectively. For this experiment, 2% transmittance and 250 ms exposure time are used in the A594 channel, and 10% transmittance and 500 ms exposure time are used for brightfield. 
		NOTE: The exposure time and percent transmittance will vary with different microscopes. To avoid overexposure, use the lowest percent transmittance and shortest exposure time possible that allows for proper visualization of the protein of interest.</li>
		<li>Under the Time-lapse tab, select the box next to Time-lapse. In the table that appears, enter 10 under the min column in the time-lapse row and 10 under the hours column in the total time row. This will run a time course taking images every 10 min for 10 h.</li>
		<li>Select the box next to Maintain Focus with Ultimate Focus to prevent stage drift during the movie. Under the Points tab, select the box next to Visit Point List.</li>
		<li>In the main menu, click on View &#62; Point List. A window called point list will pop up. Move the stage to an area of the chamber that shows a monolayer of cells. Click on Point Mark in the point list window.</li>
		<li>Move the stage to select 25-30 points without any overlap to avoid overexposing the cells. Each field will be imaged during each time course.</li>
		<li>In the point list window, select Calibrate All to set ultimate focus for each point. Under the Design tab in the Desgin/Run Experiment window, enter the range of point values (obtained from the point list window) in the box next to Visit Point List. Under the Run tab, save the file to the appropriate destination on the computer. On the Design/Run Experiment window, select the Play button (green triangle icon) to start the movie.</li>
	</ol>
	</li>
	<li>Optional method: Setting up a movie on a microscope 
	NOTE: These instructions are for an inverted microscope fitted with a slide holder that can accommodate a 24 mm x 50 mm coverslip. See Table of Materials for specifications about the microscope, camera, and imaging software used. A 60x oil-immersion objective is used for image acquisition.
	<ol>
		<li>Fit the coverslip inside the slide holder. Open the image acquisition software. Use course and fine adjustment knobs to focus cells using DIC or brightfield. Right click in the open window of the software. A drop-down menu will appear.</li>
		<li>Click on Acquisition Controls &#62; Acquisition. Click on Application &#62; Define/Run Experiment. This will open a window labeled ND Acquisition.</li>
		<li>In the window that opens, check the box next to XY. Move the stage to the desired location and click the box under Point Name to select that location as a point. Repeat this until 25-30 points are selected without any overlap to avoid overexposing the cells. Each field will be imaged during each time course.</li>
		<li>Set percent transmittance for each fluorescent channel. Because the strains produce Htb2-mCherry, the mCherry filter is used here. Under the Acquisition window that opened in step 4.3.3, navigate to the 555 nm button. Set the percent transmittance to 5% by adjusting the sliding scale under the 555 nm button until it reads 5%.</li>
		<li>Set the exposure time for each fluorescent channel. Under the Acquisition window, select 200 ms from the Exposure drop-down menu. 
		NOTE: The exposure time and percent transmittance will vary with different microscopes. To avoid overexposure, use the lowest percent transmittance and shortest exposure time possible that allows for proper visualization of the protein of interest.</li>
		<li>Click on the box next to Z in the ND Acquisition window. Set five z-stacks of the yeast cells that are 1.2 &#181;M apart.</li>
		<li>Click on the box next to &#955;. Select the appropriate channels to be used for the experiment. For DIC, ensure that Home is selected from the drop-down menu under Z pos. For mCherry, ensure that All is selected from the drop-down menu under Z pos. This ensures that, only a single section (in the middle of the z-stack) is taken for DIC.</li>
		<li>Select the box next to Time in the ND Acquisition window. Select the box under Phase. In the drop-down menu under Interval, select 10 min. Under Duration, select 10&#160;hour (s). This will run time course such that images are taken every 10 min for 10 h.</li>
	</ol>
	</li>
</ol>
5. Analysis of chromatin segregation
<ol>
	<li>Open the Fiji software. Open one field of view at a time: for each field, open the DIC and mCherry channels.</li>
	<li>Take the maximum intensity projection of the mCherry channel to obtain a single image: click on Image &#62; Stacks &#62; Z Project, select Max Intensity from the dropdown menu.</li>
	<li>Merge the DIC and mCherry channels in one image: click on Image &#62; Color &#62; Merge Channels.</li>
	<li>Follow a single cell through meiosis. After meiosis II completion, record the number of DNA masses.</li>
</ol>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sporulation+in+the+budding+yeast+Saccharomyces+cerevisiae.">Sporulation in the budding yeast Saccharomyces cerevisiae.</a> Genetics. 189 (3), 737-765 (2011).</li><li>Robinett, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=C.+al.+In+vivo+localization+of+DNA+sequences+and+visualization+of+large-scale+chromatin+organization+using+lac+operator/repressor+recognition.">C. al. In vivo localization of DNA sequences and visualization of large-scale chromatin organization using lac operator/repressor recognition.</a> Journal of Cell Biology. 135, 1685-1700 (1996).</li><li>Straight, A. F., Belmont, A. S., Robinett, C. C., Murray, A. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GFP+tagging+of+budding+yeast+chromosomes+reveals+that+protein-protein+interactions+can+mediate+sister+chromatid+cohesion.">GFP tagging of budding yeast chromosomes reveals that protein-protein interactions can mediate sister chromatid cohesion.</a> Current Biology. 6 (12), 1599-1608 (1996).</li><li>Sym, M., Engebrecht, J. A., Roeder, G. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ZIP1+is+a+synaptonemal+complex+protein+required+for+meiotic+chromosome+synapsis.">ZIP1 is a synaptonemal complex protein required for meiotic chromosome synapsis.</a> Cell. 72 (3), 365-378 (1993).</li><li>Scherthan, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chromosome+mobility+during+meiotic+prophase+in+Saccharomyces+cerevisiae.">Chromosome mobility during meiotic prophase in Saccharomyces cerevisiae.</a> Proceedings of the National Academy of Sciences of the United States of America. 104 (43), 16934-16939 (2007).</li></ol>

The authors declare no competing financial interests.

This protocol combines the NDT80-in system to synchronize cells, the anchor away technique to deplete proteins from the nucleus, and fluorescence time-lapse microscopy to image budding yeast cells during meiosis. The NDT80-in system is a method for meiotic cell cycle synchronization that utilizes a prophase I arrest and release4,8. Although individual cells will vary slightly in the amount of time spent in each of the subsequent meiotic stages, ...

Time-lapse fluorescence microcopy is a valuable tool for studying the dynamics of meiotic chromosome segregation in budding yeast1,2. Budding yeast cells can be induced to undergo meiosis through starvation of key nutrients3. During meiosis, cells undergo one round of chromosome segregation followed by two divisions to create four meiotic products that are packaged into spores (Figure 1). Individual cells can be visualized throughout each stage of meiosis, which generates spatial and temporal data that can be easily missed by fixed-cell imaging. This protocol shows how combining time-lapse fluorescence microscopy with two previously established methods, the inducible NDT80 system (NDT80-in) and the anchor away technique, can be used to study the function of specific proteins in distinct meiotic stages.
The NDT80-in system is a powerful tool for meiotic cell cycle synchronization that relies on the inducible expression of the middle meiosis transcription factor NDT804,5. NDT80 expression is required for prophase I exit6,7. With the NDT80-in system, NDT80 is under the control of the GAL1-10 promoter in cells expressing the Gal4 transcription factor fused to an estrogen receptor (Gal4-ER)4,5. Because Gal4-ER only enters the nucleus when bound to &#946;-estradiol, NDT80-in cells arrest in prophase I in the absence of &#946;-estradiol, which allows the synchronization of cells in prophase I (Figure 1). &#946;-estradiol addition promotes the translocation of the Gal4-ER transcription factor into the nucleus, where it binds GAL1-10 to drive expression of NDT80, leading to synchronous entry into the meiotic divisions. Although time-lapse microscopy can be performed without synchronization, the advantage of using synchronization is the ability to add an inhibitor or a drug while cells are at a specific stage of meiosis.
The anchor away technique is an inducible system by which a protein can be depleted from the nucleus with the addition of rapamycin8. This technique is ideal for studying nuclear proteins during cell division in budding yeast because yeast cells undergo closed mitosis and meiosis, in which the nuclear envelope does not break down. Furthermore, this technique is very useful for proteins that have multiple functions throughout meiosis. Unlike for deletions, mutant alleles, or meiotic null alleles, the removal of a target protein from the nucleus at a specific stage does not compromise target protein activity at earlier stages, allowing for a more accurate interpretation of results. The anchor away system utilizes the shuttling of ribosomal subunits between the nucleus and cytoplasm that occurs upon ribosomal maturation8. To deplete the target protein from the nucleus, the target protein is tagged with the FKBP12-rapamycin-binding domain (FRB) in a strain in which the ribosomal subunit Rpl13A is tagged with FKBP12. Without rapamycin, FRB and FKBP12 do not interact, and the FRB-tagged protein remains in the nucleus. Upon rapamycin addition, the rapamycin forms a stable complex with FKBP12 and FRB, and the complex is shuttled out of the nucleus due to the interaction with Rpl13A (Figure 1). To prevent cell death upon rapamycin addition, cells harbor the tor1-1 mutation of the TOR1 gene. Additionally, these cells contain fpr1&#916;, a null allele of the S. cerevisiae FKBP12 protein, which prevents endogenous Fpr1 from out-competing Rpl13A-FKBP12 for FRB and rapamycin binding. The anchor away background mutations, tor1-1 and fpr1&#916;, do not affect meiotic timings or chromosome segregation2.
To demonstrate the usefulness of this technique, the kinetochore protein Ctf19 was depleted at different timepoints throughout meiosis. Ctf19 is a component of the kinetochore that is dispensable in mitosis but required for proper chromosome segregation in meiosis9,10,11,12,13. In meiosis, the kinetochore is shed in prophase I, and Ctf19 is important for kinetochore re-assembly9,14. For this protocol, cells with the NDT80-in system were synchronized, and the anchor away technique was used to deplete the target protein Ctf19 from the nucleus before and after the release from prophase I, and after meiosis I chromosome segregation (Figure 1). This protocol can be adapted to deplete other proteins of interest at any stage of meiosis and mitosis.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>&#946;-estradiol</td>
			<td>Millipore Sigma</td>
			<td>E8875</td>
			<td>Make 1mM stocks in 95% EtOH</td>
		</tr>
		<tr>
			<td>0.22 uM Threaded Bottle-top Filter</td>
			<td>Millipore Sigma</td>
			<td>S2GPT02RE</td>
			<td></td>
		</tr>
		<tr>
			<td>100% EtOH</td>
			<td>Fisher Scientific</td>
			<td>22-032-601</td>
			<td></td>
		</tr>
		<tr>
			<td>10X PBS</td>
			<td>Fisher Scientific</td>
			<td>BP399500</td>
			<td>Dilute 1:10 to use as solvent for ConA</td>
		</tr>
		<tr>
			<td>24 mm x 50 mm coverslip No. 1.5</td>
			<td>VWR North American</td>
			<td>48393241</td>
			<td></td>
		</tr>
		<tr>
			<td>25 mm x 75 mm microscope slides</td>
			<td>VWR North American</td>
			<td>48300-026</td>
			<td></td>
		</tr>
		<tr>
			<td>Adenine hemisulfate salt</td>
			<td>Millipore Sigma</td>
			<td>A9126</td>
			<td>To supplement SC, SCA, and 1% Kac</td>
		</tr>
		<tr>
			<td>Bacto Agar</td>
			<td>BD</td>
			<td>214030</td>
			<td></td>
		</tr>
		<tr>
			<td>Concanavialin A</td>
			<td>Mllipore Sigma</td>
			<td>C2010</td>
			<td>Make as 1mg/mL in 1X PBS</td>
		</tr>
		<tr>
			<td>CoolSNAP HQ2 CCD camera</td>
			<td>Photometrics</td>
			<td></td>
			<td>Used in Section 4.3</td>
		</tr>
		<tr>
			<td>D-glucose</td>
			<td>Fisher Scientific</td>
			<td>D16-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Difco Yeast Nitrogen Base w/o Amino Acids</td>
			<td>BD</td>
			<td>291920</td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethyl sulfoxide (DMSO)</td>
			<td>Millipore Sigma</td>
			<td>D5879</td>
			<td></td>
		</tr>
		<tr>
			<td>Eclipse Ti2 inverted-objective micrscope</td>
			<td>Nikon</td>
			<td></td>
			<td>Used in Section 4.4</td>
		</tr>
		<tr>
			<td>Fiji</td>
			<td>NIH</td>
			<td></td>
			<td>Download from https://fiji.sc/</td>
		</tr>
		<tr>
			<td>GE Personal DeltaVision Microscope</td>
			<td>Applied Precision</td>
			<td></td>
			<td>Used in Section 4.3</td>
		</tr>
		<tr>
			<td>L-Tryptophan&#160;</td>
			<td>Millipore Sigma</td>
			<td>T0254</td>
			<td>To supplement SC, SCA, and 1% Kac</td>
		</tr>
		<tr>
			<td>Modeling Clay</td>
			<td>Crayola</td>
			<td>&#160;2302880000</td>
			<td>To secure coverslip in slide holder</td>
		</tr>
		<tr>
			<td>NIS-Elements AR 5.30.04 Imaging Software</td>
			<td>Nikon</td>
			<td></td>
			<td>Used in Section 4.4</td>
		</tr>
		<tr>
			<td>ORCA-Fustion BT Camera</td>
			<td>Hamamatsu</td>
			<td>C15440-20UP</td>
			<td>Used in Section 4.4</td>
		</tr>
		<tr>
			<td>Plastic pipette tip holder</td>
			<td>Dot Scientific</td>
			<td>LTS1000-HR</td>
			<td>Cut a 4 square x 4 square section of the rack portion of this product.&#160;</td>
		</tr>
		<tr>
			<td>Pottassium Acetate</td>
			<td>Fisher Scientific</td>
			<td>BP264</td>
			<td></td>
		</tr>
		<tr>
			<td>Rapamycin</td>
			<td>Fisher Scientific</td>
			<td>BP29631</td>
			<td>Make 1mg/mL stocks in DMSO</td>
		</tr>
		<tr>
			<td>Silicone Sealant</td>
			<td>Aqueon</td>
			<td>100165001</td>
			<td>Also known as aquarium glue.</td>
		</tr>
		<tr>
			<td>SoftWorx7.0.0&#160; Imaging Software</td>
			<td>Applied Precision</td>
			<td></td>
			<td>Used in Section 4.3</td>
		</tr>
		<tr>
			<td>Synthetic Complete Mixture (Kaiser)&#160;</td>
			<td>Formedium</td>
			<td>DSCK2500</td>
			<td></td>
		</tr>
		<tr>
			<td>Type N immersion oil&#160;</td>
			<td>Nikon</td>
			<td>MXA22166</td>
			<td></td>
		</tr>
	</tbody>
</table>

use of time-lapse microscopy and stage-specific nuclear depletion of proteins to study meiosis in s. cerevisiae

Time-lapse fluorescence microscopy has revolutionized the understanding of meiotic cell-cycle events by providing temporal and spatial data that is often not seen by imaging fixed cells. Budding yeast has proved to be an important model organism to study meiotic chromosome segregation because many meiotic genes are highly conserved. Time-lapse microscopy of meiosis in budding yeast allows the monitoring of different meiotic mutants to show how the mutation disrupts meiotic processes. However, many proteins function at multiple points in meiosis. The use of loss-of-function or meiotic null mutants can therefore disrupt an early process, blocking or disturbing the later process and making it difficult to determine the phenotypes associated with each individual role. To circumvent this challenge, this protocol describes how the proteins can be conditionally depleted from the nucleus at specific stages of meiosis while monitoring meiotic events using time-lapse microscopy. Specifically, this protocol describes how the cells are synchronized in prophase I, how the anchor away technique is used to deplete proteins from the nucleus at specific meiotic stages, and how time-lapse imaging is used to monitor meiotic chromosome segregation. As an example of the usefulness of the technique, the kinetochore protein Ctf19 was depleted from the nucleus at different time points during meiosis, and the number of chromatin masses was analyzed at the end of meiosis II. Overall, this protocol can be adapted to deplete different nuclear proteins from the nucleus while monitoring the meiotic divisions.

To monitor chromatin segregation, histone protein Htb2 was tagged with mCherry. In prophase I, the chromatin appears as a single Htb2 mass. After homologous chromosomes segregate in the first meiotic division, the chromatin appears as two distinct masses (Figure 3A). After the sister chromatids segregate, the chromatin appears as four masses. If some chromosomes fail to attach to spindle microtubules, additional masses can be seen after meiosis I or meiosis II.
Th...

Watch this Scientific Journal Video about Use of Time-Lapse Microscopy and Stage-Specific Nuclear Depletion of Proteins to Study Meiosis in S. cerevisiae at JoVE.com

Use of Time-Lapse Microscopy and Stage-Specific Nuclear Depletion of Proteins to Study Meiosis in S. cerevisiae

Time-lapse microscopy is a valuable tool for studying meiosis in budding yeast. This protocol describes a method that combines cell-cycle synchronization, time-lapse microscopy, and conditional depletion of a target protein to demonstrate how to study the function of a specific protein during meiotic chromosome segregation.

Use of Time-Lapse Microscopy and Stage-Specific Nuclear Depletion of Proteins to Study Meiosis in S. cerevisiae

Time-lapse fluorescence microscopy has revolutionized the understanding of meiotic cell-cycle events by providing temporal and spatial data that is often ...

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1.7K Views.  Indiana University. Time-lapse microscopy is a valuable tool for studying meiosis in budding yeast. This protocol describes a method that combines cell-cycle synchronization, time-lapse microscopy, and conditional depletion of a target protein to demonstrate how to study the function of a specific protein during meiotic chromosome segregation.

Video: Use of Time-Lapse Microscopy and Stage-Specific Nuclear Depletion of Proteins to Study Meiosis in S. cerevisiae

Assay for Adhesion and Agar Invasion in S. cerevisiae

Yeasts are found in natural biofilms, where many microorganisms colonize surfaces. In artificial environments, such as surfaces of man-made objects, biofilms can reduce industrial productivity, destroy structures, and threaten human life. 1-3 On the other hand, harnessing the power of biofilms can help clean the environment and generate sustainable energy. 4-8
The ability of S. cerevisiae to colonize surfaces and participate in complex biofilms was mostly ignored until the rediscovery of the differentiation programs triggered by various signaling pathways and environmental cues in this organism. 9, 10 The continuing interest in using S. cerevisiae as a model organism to understand the interaction and convergence of signaling pathways, such as the Ras-PKA, Kss1 MAPK, and Hog1 osmolarity pathways, quickly placed S. cerevisiae in the junction of biofilm biology and signal transduction research. 11-20 To this end, differentiation of yeast cells into long, adhesive, pseudohyphal filaments became a convenient readout for the activation of signal transduction pathways upon various environmental changes. However, filamentation is a complex collection of phenotypes, which makes assaying for it as if it were a simple phenotype misleading. In the past decade, several assays were successfully adopted from bacterial biofilm studies to yeast research, such as MAT formation assays to measure colony spread on soft agar and crystal violet staining to quantitatively measure cell-surface adherence. 12, 21 However, there has been some confusion in assays developed to qualitatively assess the adhesive and invasive phenotypes of yeast in agar.
Here, we present a simple and reliable method for assessing the adhesive and invasive quality of yeast strains with easy-to-understand steps to isolate the adhesion assessment from invasion assessment. Our method, adopted from previous studies, 10, 16 involves growing cells in liquid media and plating on differential nutrient conditions for growth of large spots, which we then wash with water to assess adhesion and rub cells completely off the agar surface to assess invasion into the agar. We eliminate the need for streaking cells onto agar, which affects the invasion of cells into the agar. In general, we observed that haploid strains that invade agar are always adhesive, yet not all adhesive strains can invade agar medium. Our approach can be used in conjunction with other assays to carefully dissect the differentiation steps and requirements of yeast signal transduction, differentiation, quorum sensing, and biofilm formation.

We describe a qualitative assay for yeast adhesion and agar invasion as a measure of invasive and pseudohyphal differentiation. This simple assay can be used to assess the invasive phenotype of various mutants as well as the effects environmental cues and signaling pathways on yeast differentiation.

Test dla Przyczepność i Invasion Agar w S. cerevisiae

Yeasts are found in natural biofilms, where many microorganisms colonize surfaces. In artificial environments, such as surfaces of man-made objects, ...

Monitoring Actin Disassembly with Time-lapse Microscopy

Okay, so what I'd like to do today is to show you how to construct a flow chamber, a fusion chamber for imaging on an upright microscope. What then I then do is I create two spaces using film. So what I do is I take the para film, I cut it into small strips.Next I'm going to, this will basically form the channels for the, the walls, and I dry them with a tilted air. I put the cover slip on top of the para, And here's the finished. Then I heat it on a heat block to melt the param and basically create a seal.So what this is, is the, the cover glass here, and then on top of the cover glass is two layers of phim as you see right here. And then on top of this, again, there's a, it's a 22 by two two mil meter cover slip. This one right here, which I pressed against the param to fall the seal.Okay, so the liquid goes in in one direction here and I can suck it out with a filter paper here in this direction. And I'll show you how to do that in a bit. So what I like to do is I like to put it in a humidified chamber, which is very simple.Take two kind of elevated strips and have a filter of paper underneath. And to keep it noisy as I, I put the cover, cover slip on the flow chamber on. Now the way you introduce a solution to the chamber is that you just put the pipet tip at the entrance of the chamber and you just split.Now what I do is I incubate the chamber with the lid closed for 10 minutes. What I usually do to suck the liquid out from the other side is to use it, get a filter paper and cut it into thin strips like this. And I take one of these thin strips and I simultaneously apply the solution into the entrance while putting the filter paper at the exit to suck the exchange, the solution.So you'll see the liquid flowing through the solution. And then I wait for five minutes for the mid close. Now what I'm ready to do is to actually put that the actin it and the actin, I actually have to polymerize, I polymerize inside the flow chamber and it's just the same procedures before.So because like this, and then now that the action is polymerized, I wash out all the un excess un polymerized action using a large volume of just buffer. So here I have about a hundred microliters and it's a few reaction chamber volumes worth of buffer. Just, you know, when the filter paper becomes saturated, I have to change the direction.Okay?And so I'm ready to do imaging with this. So perfusion chamber with actin attached to it. Now I take the slide, mount it under the microscope, and since I'm imaging with an oil objective, I'm gonna put some immersion oil on the top surface of the cover, grass touch, such so assess solution or protein or anything on acting inside.So what I do is I can actually exchange the solution side while I'm doing the time lapse imaging. As such.

Monitoring Actin Demontaż z poklatkowy Mikroskopia

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

A high-throughput method to globally study the organelle morphology in S. cerevisiae

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help achieve an comprehensive understanding of molecular pathways. In Saccharomyces cerevisiae, with the development of the non-essential gene deletion array, we can globally study the morphology of different organelles like the endoplasmic reticulum (ER) and the mitochondria using GFP (or variant)-markers in different gene backgrounds. However, incorporating GFP markers in each single mutant individually is a labor-intensive process. Here, we describe a procedure that is routinely used in our laboratory. By using a robotic system to handle high-density yeast arrays and drug selection techniques, we can significantly shorten the time required and improve reproducibility. In brief, we cross a GFP-tagged mitochondrial marker (Apc1-GFP) to a high-density array of 4,672 nonessential gene deletion mutants by robotic replica pinning.  Through diploid selection, sporulation, germination and dual marker selection, we recover both alleles. As a result, each haploid single mutant contains Apc1-GFP incorporated at its genomic locus. Now, we can study the morphology of mitochondria in all non-essential mutant background. Using this high-throughput approach, we can conveniently study and delineate the pathways and genes involved in the inheritance and the formation of organelles in a genome-wide setting.

GFP-fusion proteins are widely used to visualize organelles by confocal microscopy. However, screening for mutations that affect the morphology of organelles generally requires individual mutagenesis and is time consuming. Here, we demonstrate a method to simultaneously incorporate organelle-GFP markers in almost 5,000 non-essential genes in yeast.

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help ...

Identification of protein complexes with quantitative proteomics in S. cerevisiae

Lipids are the building blocks of cellular membranes that function as barriers and in compartmentalization of cellular processes, and recently, as important intracellular signalling molecules. However, unlike proteins, lipids are small hydrophobic molecules that traffic primarily by poorly described nonvesicular routes, which are hypothesized to occur at membrane contact sites (MCSs). MCSs are regions where the endoplasmic reticulum (ER) makes direct physical contact with a partnering organelle, e.g., plasma membrane (PM). The ER portion of ER-PM MCSs is enriched in lipid-synthesizing enzymes, suggesting that lipid synthesis is directed to these sites and implying that MCSs are important for lipid traffic. Yeast is an ideal model to study ER-PM MCSs because of their abundance, with over 1000 contacts per cell, and their conserved nature in all eukaryotes. Uncovering the proteins that constitute MCSs is critical to understanding how lipids traffic is accomplished in cells, and how they act as signaling molecules. We have found that an ER called Scs2p localize to ER-PM MCSs and is important for their formation. We are focused on uncovering the molecular partners of Scs2p. Identification of protein complexes traditionally relies on first resolving purified protein samples by gel electrophoresis, followed by in-gel digestion of protein bands and analysis of peptides by mass spectrometry. This often limits the study to a small subset of proteins. Also, protein complexes are exposed to denaturing or non-physiological conditions during the procedure. To circumvent these problems, we have implemented a large-scale quantitative proteomics technique to extract unbiased and quantified data. We use stable isotope labeling with amino acids in cell culture (SILAC) to incorporate staple isotope nuclei in proteins in an untagged control strain. Equal volumes of tagged culture and untagged, SILAC-labeled culture are mixed together and lysed by grinding in liquid nitrogen. We then carry out an affinity purification procedure to pull down protein complexes. Finally, we precipitate the protein sample, which is ready for analysis by high-performance liquid chromatography/ tandem mass spectrometry. Most importantly, proteins in the control strain are labeled by the heavy isotope and will produce a mass/ charge shift that can be quantified against the unlabeled proteins in the bait strain. Therefore, contaminants, or unspecific binding can be easily eliminated. By using this approach, we have identified several novel proteins that localize to ER-PM MCSs. Here we present a detailed description of our approach. 

Here we describe a new quantitative proteomics technique for identifying protein complexes in Saccharomyces cerevisiae. In this study, we have used the SILAC method together with affinity purification followed by tandem mass spectrometry to identify with high specificity the binding partners of an ER protein, Scs2p.

Lipids are the building blocks of cellular membranes that function as barriers and in compartmentalization of cellular processes, and recently, as ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes. The transparency of the 
embryos, the genetic resources, and the relative ease of transformation are qualities that make C. elegans an excellent model for early embryogenesis. Laser-based 
confocal microscopy and fluorescently labeled tags allow researchers to follow specific cellular structures and proteins in the developing embryo. For example, 
one can follow specific organelles, such as lysosomes or mitochondria, using fluorescently labeled dyes. These dyes can be delivered to the early embryo by means 
of microinjection into the adult gonad. Also, the localization of specific proteins can be followed using fluorescent protein tags. Examples are presented here 
demonstrating the use of a fluorescent lysosomal dye as well as fluorescently tagged histone and ubiquitin proteins. The labeled histone is used to visualize 
the DNA and thus identify the stage of the cell cycle. GFP-tagged ubiquitin reveals the dynamics of ubiquitinated vesicles in the early embryo. Observations 
of labeled lysosomes and GFP:: ubiquitin can be used to determine if there is colocalization between ubiquitinated vesicles and lysosomes. A technique for 
the microinjection of the lysosomal dye is presented. Techniques for generating transgenenic strains are presented elsewhere (1, 2). For imaging, embryos 
are cut out of adult hermaphrodite nematodes and mounted onto 2% agarose pads followed by time-lapse microscopy on a standard laser scanning confocal 
microscope or a spinning disk confocal microscope. This methodology provides for the high resolution visualization of early embryogenesis.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

This article describes a technique for the visualization of the early events of embryogenesis in the nematode Caenorhabditis elegans.

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes.  The transparency of the 
embryos, the genetic ...

Quantitative Analysis of Random Migration of Cells Using Time-lapse Video Microscopy

Cell migration is a dynamic process, which is important for embryonic development, tissue repair, immune system function, and tumor invasion 1, 2. During directional migration, cells move rapidly in response to an extracellular chemotactic signal, or in response to intrinsic cues 3 provided by the basic motility machinery. Random migration occurs when a cell possesses low intrinsic directionality, allowing the cells to explore their local environment.
Cell migration is a complex process, in the initial response cell undergoes polarization and extends protrusions in the direction of migration 2. Traditional methods to measure migration such as the Boyden chamber migration assay is an easy method to measure chemotaxis in vitro, which allows measuring migration as an end point result. However, this approach neither allows measurement of individual migration parameters, nor does it allow to visualization of morphological changes that cell undergoes during migration.
Here, we present a method that allows us to monitor migrating cells in real time using video - time lapse microscopy. Since cell migration and invasion are hallmarks of cancer, this method will be applicable in studying cancer cell migration and invasion in vitro. Random migration of platelets has been considered as one of the parameters of platelet function 4, hence this method could also be helpful in studying platelet functions. This assay has the advantage of being rapid, reliable, reproducible, and does not require optimization of cell numbers. In order to maintain physiologically suitable conditions for cells, the microscope is equipped with CO2 supply and temperature thermostat. Cell movement is monitored by taking pictures using a camera fitted to the microscope at regular intervals. Cell migration can be calculated by measuring average speed and average displacement, which is calculated by Slidebook software.

This method allows monitoring of cells in real time and quantitative measurements of different cell migration parameters such as speed, displacement, and velocity. Unlike the traditional methods, this real time approach is not based on endpoint quantitative migration measurements; instead it allows monitoring and calculating different parameters continuously.

Cell migration is a dynamic process, which is important for embryonic development, tissue repair, immune system function, and tumor invasion 1, 2. During ...

Flow Cytometry-based Purification of S. cerevisiae Zygotes

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae zygotes result from the fusion of haploid cells of distinct mating type (MATa, MATalpha) and give rise to corresponding stable diploids that successively generate as many as 20 diploid progeny as a result of their strikingly asymmetric mitotic divisions 2. Zygote formation is orchestrated by a complex sequence of events: In this process, soluble mating factors bind to cognate receptors, triggering receptor-mediated signaling cascades that facilitate interruption of the cell cycle and culminate in cell-cell fusion. Zygotes may be considered a model for progenitor or stem cell function.
Although much has been learned about the formation of zygotes and although zygotes have been used to investigate cell-molecular questions of general significance, almost all studies have made use of mating mixtures in which zygotes are intermixed with a majority population of haploid cells 3-8. Many aspects of the biochemistry of zygote formation and the continuing life of the zygote therefore remain uninvestigated.
Reports of purification of yeast zygotes describe protocols based on their sedimentation properties 9; however, this sedimentation-based procedure did not yield nearly 90% purity in our hands. Moreover, it has the disadvantage that cells are exposed to hypertonic sorbitol. We therefore have developed a versatile purification procedure. For this purpose, pairs of haploid cells expressing red or green fluorescent proteins were co-incubated to allow zygote formation, harvested at various times, and the resulting zygotes were purified using a flow cytometry-based sorting protocol. This technique provides a convenient visual assessment of purity and maturation. The average purity of the fraction is approximately 90%. According to the timing of harvest, zygotes of varying degrees of maturity can be recovered. The purified samples provide a convenient point of departure for "-omic" studies, for recovery of initial progeny, and for systematic investigation of this progenitor cell.

Flow Cytometry-based Purification of S. cerevisiae Zygotes

To purify zygotes of S. cerevisiae, haploid cells of opposite mating type were engineered to express red or green fluorescent proteins, co-incubated to allow zygote formation, and fractionated using a flow cytometry-based protocol. The highly-enriched fraction enables subsequent "-omic" studies, recovery of initial progeny, and systematic investigation of zygote morphogenesis.

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae ...

Use of Time Lapse Microscopy to Visualize Anoxia-induced Suspended Animation in C. elegans Embryos

Caenorhabdits elegans has been used extensively in the study of stress resistance, which is facilitated by the transparency of the adult and embryo stages as well as by the availability of genetic mutants and transgenic strains expressing a myriad of fusion proteins1-4. In addition, dynamic processes such as cell division can be viewed using fluorescently labeled reporter proteins. The study of mitosis can be facilitated through the use of time-lapse experiments in various systems including intact organisms; thus the early C. elegans embryo is well suited for this study. Presented here is a technique by which in vivo imaging of sub-cellular structures in response to anoxic (99.999% N2; &lt;2 ppm O2) stress is possible using a simple gas flow through setup on a high-powered microscope. A microincubation chamber is used in conjunction with nitrogen gas flow through and a spinning disc confocal microscope to create a controlled environment in which animals can be imaged in vivo. Using GFP-tagged gamma tubulin and histone, the dynamics and arrest of cell division can be monitored before, during and after exposure to an oxygen-deprived environment. The results of this technique are high resolution, detailed videos and images of cellular structures within blastomeres of embryos exposed to oxygen deprivation.

Use of Time Lapse Microscopy to Visualize Anoxia-induced Suspended Animation in C. elegans Embryos

Described here is an in vivo technique to image sub-cellular structures in animals exposed to anoxia using a gas flow through microincubation chamber in conjunction with a spinning disc confocal microscope. This method is straightforward and flexible enough to suit a variety of experimental parameters and model systems.

Caenorhabdits elegans has been used extensively in the study of stress resistance, which is facilitated by the transparency of the adult and embryo stages ...

Use of Time-Lapse Microscopy and Stage-Specific Nuclear Depletion of Proteins to Study Meiosis in S. cerevisiae

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Time-lapse Imaging of Mitosis After siRNA Transfection

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Quantitative Analysis of Random Migration of Cells Using Time-lapse Video Microscopy

Flow Cytometry-based Purification of S. cerevisiae Zygotes

Use of Time Lapse Microscopy to Visualize Anoxia-induced Suspended Animation in C. elegans Embryos