This work was supported by startup funds from The University of Tennessee and TN-SCORE, a multi-disciplinary research program sponsored by NSF-EPSCoR (EPS-1004083).

1. Preparation of Sample
<ol>
	<li>Grow fission yeast cells expressing the spindle pole body marker Sad1-mCherry23 and type II myosin regulatory light chain Rlc1-Tomato24 in YE liquid media at 32 &#176;C for 8 generations. 
	NOTE: For temperature sensitive mutants, grow cells at 25 &#176;C. Grow cells to mid-log phase to an O.D.600 of 0.5 in YE. For more on fission yeast growth conditions refer to the Fission yeast lab manual25.</li>
	<li>Prepare YE (or minimal) media with 0.6% agarose (Agarose based media show less fluorescent background in comparison to agar). To the melted media add 1 mM of ascorbic acid (vitamin C). The vitamin C quenches free radicals that are released due to photo-bleaching, thus minimizing photo-toxicity.</li>
	<li>Pour 3 - 4 mL of the vitamin C laced melted media on to a glass bottomed culture dish. Let the media cool and solidify. 
	NOTE: The thickness of the glass in the culture dish depends on the cover slip thickness suitable for the microscope. Here, use coverslip No. 1.5.</li>
	<li>Gently raise the solidified media from the culture dish with the help of a sharp scalpel. Place a pipette tip between the media slab and the culture dish to prevent the slab from falling back into the dish (Figure 1).</li>
	<li>Take 1 mL of freshly growing cells as described in 1.1 above. Spin the cells gently at 300 x g for 30 s. Discard the supernatant. Resuspend the cells in the small amount of residual media that remains in the tube after discarding the supernatant.</li>
	<li>Load 2 - 5 &#181;L of resuspended cell culture between the media slab and the glass bottom of the culture dish. The cells should be on the glass. The pipette tip placed between the media and the dish enables easy loading of the cell culture.</li>
	<li>Very gently remove the pipette tip and place the media slab back in its original position on the culture dish. Make sure to avoid any air bubbles. If need be gently press out the air bubbles by sliding the back of a pipette tip on top of the media slab.</li>
	<li>Let the cells sit in the culture dish for 30 min&#160;to one hour in the dark at the temperature at which microscopy will be performed. This is important to prevent any shock due to change in growth conditions for the cells.</li>
</ol>
2. Image Acquisition
<ol>
	<li>Place a drop of oil on the outer side of the glass on the culture dish. Place the culture dish on an inverted microscope.</li>
	<li>To minimize auto fluorescence of YE media use a GFP filter with a narrow wavelength range (520 - 535 nm).</li>
	<li>Focus on the medial plane of the cells and ensure that they are well spaced and not too crowded. Make sure to start image acquisition of the cells in the suitable stage of the cell cycle. For this experiment, follow cells from late G2.</li>
	<li>Program the image acquisition software to take GFP, RFP and DIC images of the cells.
	<ol>
		<li>For this experiment use 100 ms exposure time for DIC and 75 ms exposure time for GFP and RFP filters. The exposure time depends on the settings of the microscope, the protein under study and the duration of the experiment. For the given microscope settings, use 50% laser power.</li>
		<li>To capture images in 3 dimensions, program the image acquisition software for Z-scale imaging. Use a step size of 0.4 &#181;m and a distance of 3.0 &#181;m (half of total z) around the central focus point of the cells. This leads to the capture of 16 Z-frames per time point with a total Z distance of 6 &#181;m. 
		NOTE: Z-series image acquisition allows the capture of the spindle pole bodies.</li>
	</ol>
	</li>
	<li>Take images every 2 min. Change the exposure time according to the requirements of the experiment. The exposure time can also be changed according to the protein of interest and signal strength. However, note that shorter intervals or longer exposure time increase photo-toxicity due to bleaching and hence cells can be followed only for a short time.</li>
	<li>Acquire images till the cells physically separate as observed by the DIC images, or program the software to stop acquisition after 90 min.</li>
</ol>
3. Image Analysis
<ol>
	<li>Temporal analysis of cytokinetic events
	<ol>
		<li>Using the image acquisition software make projections of the Z-frames for each time-point. Make different files for each wavelength of image acquired.</li>
		<li>Export this projection time-point series as a .tiff file.</li>
		<li>Analyze the images using ImageJ software. Open the image series for any one of the wavelengths.</li>
		<li>Select a cell to be studied. Double click on the line option of toolbar. This will open another window to adjust line width. Adjust the line width to correspond to the cell width. For a WT cell this is about 40 - 50 pixels. Draw a line along the long axis of the cell of interest.</li>
		<li>Click on Analyze&#62;Tools&#62;ROI manager and click on add. This will add the selected line to the ROI window for use later. Do not close this window.</li>
		<li>Click on the image of interest and select Edit&#62;Selection&#62;Straighten. This will open another window. Check &#34;Process entire stack&#34;. This will straighten the cell horizontally.</li>
		<li>Click on Image&#62;Transform&#62;Rotate 90 Degrees Right. This image is now straightened vertically.</li>
		<li>Click on Image&#62;Stacks&#62;Make Montage. This will open a window to assign the number of rows and columns for the stack, the scale factor for the image size, the first and last slice (frame) for the montage and the frame increment for the montage. Check label slices and hit enter.</li>
		<li>As a window with a montage of the cell of interest is shown, open the image series for the other wavelength.</li>
		<li>Click on the line identifier on ROI manager. This will select the same cell on the second image file. Repeat steps 3.1.6 - 3.1.8. This will open a montage of the second image.</li>
		<li>On the image with the spindle pole body marker Sad1-mCherry, look for onset of separation of the spindle pole body marker and mark that time point as time 0.</li>
		<li>Follow the Rlc1-Tomato signal on the image over time and look for the signal that appears as a distinct line as opposed to patches of Rlc1-Tomato. This time-point marks completion of actomyosin ring assembly/start of maturation phase.</li>
		<li>Next scroll through the movie over time to determine when the Rlc1-Tomato ring starts to decrease in size. This is marked as the end of maturation phase/onset of ring constriction.</li>
		<li>Follow the Rlc1-Tomato signal over time throughout constriction until it appears as a dot in the middle of the cell axis. This is marked as the end of ring constriction/end of septum ingression.</li>
		<li>Following the montage of DIC images, determine the final cell separation. Record the time point when the cell physically separates.</li>
		<li>For spindle pole body separation measure the distance between the two spindle pole bodies over time using the line tool in ImageJ. Plot the distance in spindle pole body over time.</li>
		<li>Record the different time-points for the different cytokinetic events to calculate the duration of each cytokinetic phase and compare with spindle pole body separation over time.</li>
	</ol>
	</li>
	<li>Spatial analysis of cytokinetic events
	<ol>
		<li>Select a cell with a visible actomyosin ring. Double click on the line option of ImageJ toolbar. This will open another window to adjust line width. Adjust the line width to correspond to the ring thickness (15 - 20 pixels). Draw a line along the ring taking care to ensure the entire thickness of the ring is included.</li>
		<li>Click on Analyze&#62;Tools&#62;ROI manager and click on add. This will add the selected line to the ROI window for use later. Do not close this window.</li>
		<li>Click on the image of interest and select Edit&#62;Selection&#62;Straighten. This will open another window. Check &#34;Process entire stack&#34;. This will straighten the ring horizontally.</li>
		<li>Reset the intensity of the brightest plane in the straightened z-stack (from 3.1.6) using Image&#62;Adjust&#62;Brightness/Contrast&#62;Reset.</li>
		<li>Click on the tab Stk&#62;3D Project. This will open a dialog box. Change the projection method to Brightest Point and the Slice spacing to 2 or 3 pixels. Finally, change axis of rotation to X or Y-axis (this will change depending upon the desired viewing angle), click interpolate and click OK to generate a 3D projection of the image. Use the scroll bar beneath the image to rotate the image.</li>
		<li>Open the image series for the other wavelength using ImageJ.</li>
		<li>Click on the line identifier on ROI manager. This will select the same ring on the second image file. Repeat steps 4.2.4 and 4.2.5. This will open a 3 dimensional ring in the image with the other wavelength.</li>
		<li>With the both 3D image rings open, click on Image&#62;Color&#62;Merge Channels. This will open a box. Select each image from the drop down menu for the corresponding color desired and click OK. This will open a composite of both the images at different wavelengths.</li>
		<li>Compare the localization of different markers with regards to localization at the cytokinetic ring or ingressing membrane. Rlc1-GFP is a marker for the cytokinetic ring. Proteins co-localizing with Rlc1-GFP localize to the ring while proteins localizing behind Rlc1-GFP signal localize to the ingressing membrane. 
		NOTE: The protein of interest needs to be of a different fluorophore than that of the ring marker.</li>
		<li>Generate 3 dimensional rings during different stages of cytokinesis to compare the spatial localization of the proteins all through cytokinesis.</li>
	</ol>
	</li>
	<li>Analysis of protein distribution at cytokinetic ring
	<ol>
		<li>To compare and analyze distribution of proteins along the ring measure the co-efficient of variance of protein distribution. A high co-efficient of variance indicates uneven distribution of proteins along the division site and suggests improper ring assembly, maturation or septum formation, depending on which phase is imaged or analyzed.</li>
		<li>Using 3 dimensional reconstructed cytokinetic ring (from 3.2.2), divide the image into at least 4 quadrants such that a quarter of the ring appears in each quadrant. For this use the line tool to draw perpendicular lines on the image. After each line click Image&#62;Overlay&#62;Add selection or use short cut Ctrl+B.</li>
		<li>Go to Analyze&#62;Set measurement. Select mean gray value in the box that opens. Click OK.</li>
		<li>Using the lines drawn above as guides draw a rectangle to define each quadrant using the rectangle tool.</li>
		<li>To measure the Mean intensity of each quadrant, click on Analyze&#62;Measure or use the short cut Ctrl+M.</li>
		<li>Record the mean gray value for all four quadrants (MG1-MG4).</li>
		<li>Select a small area in each quadrant outside the ring as background selection.</li>
		<li>Measure mean gray value for Background selection for all four quadrants (BG1-BG4).</li>
		<li>li&#62;For background subtraction for each quadrant use the following formula- MG1-Average(BG1:BG4)= Intensity of ring in each quadrant (IRQ). At this stage there are 4 IRQ values, one for each quadrant.</li>
		<li>Next calculate the coefficient of variance for each ring using the following formula- Standard deviation (IRQ1:IRQ4)/Average(IRQ1:IRQ4). Calculate average Co-efficient of Variance for each strain and compare to the other strains. 
		NOTE: A statistically significant increase in co-efficient of variance in a strain compared to controls indicates uneven protein distribution/delivery along the division site at that stage of cytokinesis.</li>
	</ol>
	</li>
</ol>

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The authors declare no competing financial interests.

Here we have described a protocol to study cytokinetic events in fission yeast in a temporal manner. The protocol described here provides temporal resolution of different cytokinetic events with reference to each other; timing of protein recruitment or loss with reference to cytokinetic phase; structure of the ring throughout the different phases of cytokinesis; and the progression of cytokinesis with reference to mitosis. With this protocol it is possible to precisely define the cytokinetic phase that may be altered in ...

Cytokinesis, the final step in cell division, is a complex process essential for proper differentiation, development, and survival of an organism. Cytokinesis involves multiple events that are organized to ensure successful cell separation while maintaining genomic integrity1. Cytokinesis involves events where an actomyosin ring once assembled undergoes constriction, which is concurrent with membrane expansion and furrowing, and extracellular cellular matrix remodeling, finally followed by cell abscission1,2,3. Improper organization of cytokinetic events can lead to cell separation and ploidy defects, and may cause diseases like cancer4,5,6,7,8. The fundamental principles that enable organization of cytokinetic events are not well understood, thus leading to roadblocks in our understanding of the etiology of these diseases.
The fission yeast Schizosaccharomyces pombe (S.&#160;pombe)&#160;is an excellent model system to study cytokinesis due to the conserved nature of the proteins involved1. In fission yeast, after actomyosin ring assembly, the ring enters a maturation/dwell phase where it does not constrict9. Maturation ends with initiation of actomyosin ring constriction, concurrent with membrane furrowing and septum ingression. Seminal work over the years have given us a fairly good understanding of actomyosin ring assembly in fission yeast1,9,10. In some eukaryotes, including fission yeast, successful assembly of the actomyosin ring is not sufficient for membrane furrowing. In fission yeast, ring constriction alone does not provide sufficient force to overcome internal turgor pressure for furrow formation11. A recent model indicates that this force is instead provided by septum ingression11. In another model, the role of plasma membrane extension has been suggested to contribute to furrow formation12,13. Ring constriction and membrane furrowing do not occur in Bgs1/Cps1 temperature sensitive mutant cps1-191, the major enzyme for primary septum formation14,15. Cells lacking Bgs1 show defective primary septum and prolonged ring constriction15,16. Bgs1 is recruited to the cell division site for septum ingression during maturation after actomyosin ring assembly17,18. Similarly, during cellularization in Drosophila embryos, ring constriction is biphasic with a significantly slow initial constriction rate19, resembling the maturation phase observed in fission yeast. Biphasic ring constriction may slow down membrane furrowing to allow for sufficient membrane expansion20 and modification of extracellular matrix. This suggests that after actomyosin ring assembly, ring constriction occurs efficiently only when the cell has satisfied the requirements for furrow formation. It is not well understood what conditions are required for actomyosin ring constriction after ring assembly, nor the molecular events that regulate this process. We have recently shown that following actomyosin ring assembly the small GTPase Cdc42 undergoes a unique spatiotemporal activation pattern21. This pattern is established by the unique localization pattern of the Cdc42 guanidine nucleotide exchange factors (GEFs) that activate Cdc42. The GEF Gef1 localizes to the assembled actomyosin ring and promotes onset of ring constriction and septum ingression, while Scd1 localizes to the furrowing membrane and promotes normal septum formation. We find that the Cdc42 activation pattern established by its GEFs lead to the regulation of distinct cytokinetic events.
To understand the molecular mechanism of events that eventually lead to cell separation following actomyosin ring assembly, one needs to follow the distinct cytokinetic events in time and space. In fission yeast, cytokinesis first involves the assembly of precursor nodes around the nucleus, which eventually recruit the type II myosin, the formin Cdc12, and other proteins required for actomyosin ring assembly. To time the distinct cytokinetic events and provide a frame of reference, the separation of spindle pole body markers (spindle formation) is considered as time 022. The assembly of the actomyosin ring can be followed by monitoring over time the intensity of a fluorescently tagged actomyosin ring protein such as the type II myosin regulatory light chain Rlc1. Here we describe a microscopic approach to analyze different stages of cytokinesis over time.

<table><tbody><tr><td>Yeast extract media</td><td>Sunrise Science Products</td><td>YES 225</td><td>0.5% w/v yeast extract, 3% w/v glucose, 225mg/L adenine, histidine, leucine, uracil, and lysine</td></tr><tr><td>Agarose</td><td>SeaKem LE agarose, Lonza</td><td>50001</td><td></td></tr><tr><td>Ascorbic acid</td><td>Sigma-Aldrich</td><td>A4544</td><td></td></tr><tr><td>Glass Bottomed culture dish</td><td>MatTek Corporation</td><td>P35G-1.5-14-C</td><td>Coverslip No. 1.5 was used. This will vary as per the microscope specifications used.&amp;nbsp;</td></tr><tr><td>VT-Hawk 2D array laser scanning confocal microscopy system</td><td>Visitech International, UK</td><td></td><td>with an Olympus IX-83 inverted microscope with a 100X / numerical aperture 1.49 UAPO lens (Olympus) and EM-CCD digital camera (Hamamatsu).&amp;nbsp;</td></tr><tr><td>ImageJ</td><td>NIH</td><td></td><td>Image analysis software</td></tr></tbody></table>

spatiotemporal analysis of cytokinetic events in fission yeast

Cytokinesis, the final step in cell division is critical for maintaining genome integrity. Proper cytokinesis is important for cell differentiation and development. Cytokinesis involves a series of events that are well coordinated in time and space. Cytokinesis involves the formation of an actomyosin ring at the division site, followed by ring constriction, membrane furrow formation and extra cellular matrix remodeling. The fission yeast, Schizosaccharomyces pombe (S.&#160;pombe)&#160;is a well-studied model system that has revealed with substantial clarity the initial events in cytokinesis. However, we do not understand clearly how different cytokinetic events are coordinated spatiotemporally. To determine this, one needs to analyze the different cytokinetic events in great details in both time and in space. Here we describe a microscopy approach to examine different cytokinetic events in live cells. With this approach it is possible to time different cytokinetic events and determine the time of recruitment of different proteins during cytokinesis. In addition, we describe protocols to compare protein localization, and distribution at the site of cell division. This is a basic protocol to study cytokinesis in fission yeast and can also be used for other yeasts and fungal systems.

Fission yeast cells expressing the ring marker, Rlc1-GFP (green, Figure 2) and spindle pole body marker Sad1-mCherry (red, Figure 2) were imaged during cytokinesis. Onset of spindle pole body marker (white arrow, Figures 2A, B) is considered as time 0. Rlc1-GFP signal appears at time -4 min with reference to spindle pole body separation (red arrow, Figures 2A, 2B). Rlc1-GFP signal forms a continuous ring 10 min post spindle pole body sepa...

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Spatiotemporal Analysis of Cytokinetic Events in Fission Yeast

The fission yeast, Schizosaccharomyces pombe&#160;is an excellent model system to study cytokinesis, the final stage in cell division. Here we describe a microscopy approach to analyze different cytokinetic events in live fission yeast cells.

Cytokinesis, the final step in cell division is critical for maintaining genome integrity. Proper cytokinesis is important for cell differentiation and ...

spatiotemporal-analysis-of-cytokinetic-events-in-fission-yeast

Research

JoVE Journal

Biology

7.3K Views.  University of Tennessee. The fission yeast, Schizosaccharomyces pombe is an excellent model system to study cytokinesis, the final stage in cell division. Here we describe a microscopy approach to analyze different cytokinetic events in live fission yeast cells.

Video: Spatiotemporal Analysis of Cytokinetic Events in Fission Yeast

Measuring Replicative Life Span in the Budding Yeast

Aging is a degenerative process characterized by a progressive deterioration of cellular components and organelles resulting in mortality. The budding yeast Saccharomyces cerevisiae has been used extensively to study the biology of aging, and several determinants of yeast longevity have been shown to be conserved in multicellular eukaryotes, including worms, flies, and mice 1. Due to the lack of easily quantified age-associated phenotypes, aging in yeast has been assayed almost exclusively by measuring the life span of cells in different contexts, with two different life span paradigms in common usage 2. Chronological life span refers to the length of time that a mother cell can survive in a non-dividing, quiescence-like state, and is proposed to serve as a model for aging of post-mitotic cells in multicellular eukaryotes. Replicative life span, in contrast, refers the number of daughter cells produced by a mother cell prior to senescence, and is thought to provide a model of aging in mitotically active cells. Here we present a generalized protocol for measuring the replicative life span of budding yeast mother cells. The goal of the replicative life span assay is to determine how many times each mother cell buds. The mother and daughter cells can be easily differentiated by an experienced researcher using a standard light microscope (total magnification 160X), such as the Zeiss Axioscope 40 or another comparable model. Physical separation of daughter cells from mother cells is achieved using a manual micromanipulator equipped with a fiber-optic needle. Typical laboratory yeast strains produce 20-30 daughter cells per mother and one life span experiment requires 2-3 weeks.

In this article we present a general protocol for measuring the replicative life span of yeast mother cells.

Aging is a degenerative process characterized by a progressive deterioration of cellular components and organelles resulting in mortality. The budding ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

Acquiring Fluorescence Time-lapse Movies of Budding Yeast and Analyzing Single-cell Dynamics using GRAFTS

Fluorescence time-lapse microscopy has become a powerful tool in the study of many biological processes at the single-cell level. In particular, movies depicting the temporal dependence of gene expression provide insight into the dynamics of its regulation; however, there are many technical challenges to obtaining and analyzing fluorescence movies of single cells. We describe here a simple protocol using a commercially available microfluidic culture device to generate such data, and a MATLAB-based, graphical user interface (GUI) -based software package to quantify the fluorescence images. The software segments and tracks cells, enables the user to visually curate errors in the data, and automatically assigns lineage and division times. The GUI further analyzes the time series to produce whole cell traces as well as their first and second time derivatives. While the software was designed for S. cerevisiae, its modularity and versatility should allow it to serve as a platform for studying other cell types with few modifications.

We present a simple protocol to obtain fluorescence microscopy movies of growing yeast cells, and a GUI-based software package to extract single-cell time series data. The analysis includes automated lineage and division time assignment integrated with visual inspection and manual curation of tracked data.

Fluorescence time-lapse microscopy has become a powerful tool in the study of many biological processes at the single-cell level. In particular, movies ...

Temporal Quantification of MAPK Induced Expression in Single Yeast Cells

The quantification of gene expression at the single cell level uncovers novel regulatory mechanisms obscured in measurements performed at the population level. Two methods based on microscopy and flow cytometry are presented to demonstrate how such data can be acquired. The expression of a fluorescent reporter induced upon activation of the high osmolarity glycerol MAPK pathway in yeast is used as an example. The specific advantages of each method are highlighted. Flow cytometry measures a large number of cells (10,000) and provides a direct measure of the dynamics of protein expression independent of the slow maturation kinetics of the fluorescent protein. Imaging of living cells by microscopy is by contrast limited to the measurement of the matured form of the reporter in fewer cells. However, the data sets generated by this technique can be extremely rich thanks to the combinations of multiple reporters and to the spatial and temporal information obtained from individual cells. The combination of these two measurement methods can deliver new insights on the regulation of protein expression by signaling pathways.

Two complementary methods based on flow cytometry and microscopy are presented which enable the quantification, at the single cell level, of the dynamics of gene expression induced by the activation of a MAPK pathway in yeast.

The quantification of gene expression at the  single cell level uncovers novel regulatory mechanisms obscured in measurements  performed at the population ...

Microscopy of Fission Yeast Sexual Lifecycle

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the mechanics of cell division and cell polarity. Furthermore, classical experiments on its sexual reproduction have yielded results pivotal to current understanding of DNA recombination and meiosis. More recent analysis of fission yeast mating has raised interesting questions on extrinsic stimuli response mechanisms, polarized cell growth and cell-cell fusion. To study these topics in detail we have developed a simple protocol for microscopy of the entire sexual lifecycle. The method described here is easily adjusted to study specific mating stages. Briefly, after being grown to exponential phase in a nitrogen-rich medium, cell cultures are shifted to a nitrogen-deprived medium for periods of time suited to the stage of the sexual lifecycle that will be explored. Cells are then mounted on custom, easily built agarose pad chambers for imaging. This approach allows cells to be monitored from the onset of mating to the final formation of spores.

We provide a reproducible basic method for the long-term microscopy of the fission yeast sexual lifecycle. With minor adjustments described, the presented protocol allows research focus on different steps of the reproductive process.

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the ...

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells regulate these processes and how genetic mutations impact regulation. The quantification of microtubule dynamics in metazoan models has a number of associated challenges, including a high microtubule density and limitations on genetic manipulations. In contrast, the budding yeast model offers advantages that overcome these challenges. This protocol describes a method to measure the dynamics of single microtubules in living yeast cells. Cells expressing fluorescently tagged tubulin are adhered to assembled slide chambers, allowing for stable time-lapse image acquisition. A detailed guide for high-speed, four-dimensional image acquisition is also provided, as well as a protocol for quantifying the properties of dynamic microtubules in confocal image stacks. This method, combined with conventional yeast genetics, provides an approach that is uniquely suited for quantitatively assessing the effects of microtubule regulators or mutations that alter the activity of tubulin subunits.

Budding yeast is an advantageous model for studying microtubule dynamics in vivo due to its powerful genetics and the simplicity of its microtubule cytoskeleton. The following protocol describes how to transform and culture yeast cells, acquire confocal microscopy images, and quantitatively analyze microtubule dynamics in living yeast cells.

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells ...

Examination of Mitotic and Meiotic Fission Yeast Nuclear Dynamics by Fluorescence Live-cell Microscopy

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does not interfere with cell physiology, and requires minimal experimental handling. The low levels of technical interference enable researchers to study cells across multiple cycles of mitosis and to observe meiosis from beginning to end. Using fluorescent tags such as Green Fluorescent Protein (GFP) and Red Fluorescent Protein (RFP), researchers can analyze different factors whose functions are important for processes like transcription, DNA replication, cohesion, and segregation. Coupled with data analysis using Fiji (a free, optimized ImageJ version), live-cell imaging offers various ways of assessing protein movement, localization, stability, and timing, as well as nuclear dynamics and chromosome segregation. However, as is the case with other microscopy methods, live-cell imaging is limited by the intrinsic properties of light, which put a limit to the resolution power at high magnifications, and is also sensitive to photobleaching or phototoxicity at high wavelength frequencies. However, with some care, investigators can bypass these physical limitations by carefully choosing the right conditions, strains, and fluorescent markers to allow for the appropriate visualization of mitotic and meiotic events.

Here, we present live-cell imaging which is a non-toxic microscopy method that allows researchers to study protein behavior and nuclear dynamics in living fission yeast cells during mitosis and meiosis.

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does ...

Live Cell Imaging to Assess the Dynamics of Metaphase Timing and Cell Fate Following Mitotic Spindle Perturbations

Live cell time-lapse imaging is an important tool in cell biology that provides insight into cellular processes that might otherwise be overlooked, misunderstood, or misinterpreted by the fixed-cell analysis. While the fixed cell imaging and analysis is robust and sufficient to observe cellular steady-state, it can be limited in defining a temporal order of events at the cellular level and is ill-equipped to assess the transient nature of dynamic processes including mitotic progression. In contrast, live cell imaging is an eloquent tool that can be used to observe cellular processes at the single-cell level over time and has the capacity to capture the dynamics of processes that would otherwise be poorly represented in fixed cell imaging. Here we describe an approach to generate cells carrying fluorescently labeled markers of chromatin and microtubules and their use in live cell imaging approaches to monitor metaphase chromosome alignment and mitotic exit. We describe imaging-based techniques to assess the dynamics of spindle formation and mitotic progression, including the identification of cells at various stages in mitosis, identification&#160;and tracking of mitotic defects, and analysis of spindle dynamics and mitotic cell fate following the treatment with mitotic inhibitors.

Here we present a protocol to assess the dynamics of spindle formation and mitotic progression. Our application of time-lapse imaging enables the user to identify cells at various stages of mitosis, track and identify mitotic defects, and analyze spindle dynamics and mitotic cell fate upon exposure to anti-mitotic drugs.

Live cell time-lapse imaging is an important tool in cell biology that provides insight into cellular processes that might otherwise be overlooked, ...

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.

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.

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

Spatiotemporal Analysis of Cytokinetic Events in Fission Yeast

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Chapters in this video

Measuring Replicative Life Span in the Budding Yeast

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

Acquiring Fluorescence Time-lapse Movies of Budding Yeast and Analyzing Single-cell Dynamics using GRAFTS

Temporal Quantification of MAPK Induced Expression in Single Yeast Cells

Microscopy of Fission Yeast Sexual Lifecycle

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

Examination of Mitotic and Meiotic Fission Yeast Nuclear Dynamics by Fluorescence Live-cell Microscopy

Live Cell Imaging to Assess the Dynamics of Metaphase Timing and Cell Fate Following Mitotic Spindle Perturbations

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