We would like to thank Dr. Guo-Chiuan Hung and Dr. Bharatkumar Joshi for valuable comments that improved the manuscript.

All the experiments conducted in these studies were performed in accordance with protocols approved by the Institutional Animal Care and Use Committee at the Food and Drug Administration (FDA) research facility.
1. Isolation and Culture of Mouse Embryonic Fibroblasts (MEFs)
<ol>
	<li>Isolate mouse embryonic fibroblasts (MEFs) from a PP2A-B56&#947;- mouse strain and wild type littermates by standard protocol 7,8,9.</li>
	<li>ExpandMEFs for 3 passages, freeze and store until needed for experiments 8.</li>
</ol>
2. Culturing Mefs in 2-Well Chambered Cover Glass for Live Imaging
<ol>
	<li>Prepare 500 mL of MEF growth media containing Dulbecco Modified Eagle Medium (DMEM/F12) with 10% Fetal Bovine Serum (FBS), 1 X Penicillin/Streptomycin antibiotics, L-Glutamine (200 mM) and 1X non-essential amino acids (NEAA) in a 500-mL media bottle.</li>
	<li>Thaw vials of frozen MEFs from wild type and PP2A-B56&#947;- mice at passage 3 in pre-warmed water bath at 37 &#176;C.</li>
	<li>Transfer thawed MEFs to 15-mL tubes and slowly add dropwise 20 mL of DMEM/F12 media to the 15-mL tubes using a 10-mL pipette.</li>
	<li>Transfer MEFs along with 20 mL of DMEM/F12 growth media in T75 flasks and expand them until the cells are 70% confluent. Confluency is estimated using an inverse microscope at 4x or 10x magnification.</li>
	<li>Aspirate the growth medium using a glass pasture pipette attached to a vacuum system in the hood, add 3 mL of 0.25% trypsin/EDTA and incubate at 37 &#176;C for 5 min. Add 3 mL of DMEM/F12 growth medium to stop the reaction.</li>
	<li>Centrifuge the cells at low-speed (300 x g) for 5 min at room temperature. Carefully remove the supernatant and re-suspend the cell pellet in 1 mL of DMEM/F12 growth medium pre-warmed at 37 &#176;C in a water bath.</li>
	<li>Enumerate cells using the trypan blue exclusion method or other appropriate cell counting method 8.</li>
	<li>Seed approximately 20,000 MEFs/well in a 2-well chambered cover glass. Add 200 &#181;L/well DMEM/F12 growth medium and allow the cells to attach by incubating them overnight at 37 &#176;C and 5% CO2.</li>
</ol>
3. Synchronization
<ol>
	<li>Synchronize MEFs in G0/G1 phase by incubating cells in DMEM/F12 growth media containing 0.1% FBS for 24 h, to obtain a maximum number of cells in G0/G1 phase. 
	Note: Although serum starvation was used as a method of preference, various other methods can be used depending on the mitotic stage at which cells are needed to be arrested 10.</li>
</ol>
4. Labeling
<ol>
	<li>Prepare 1.5 mg/mL stock solution of nocodazole in DMSO. Nocodazole arrests cells at G2/M phase by inhibiting microtubule formation 10.</li>
	<li>Three days post synchronization, add 200 &#181;L of growth medium (with 10% FCS), 200 ng/mL nocodazole and CL-HB regent (Histone 2B-GFP BacMam 2.0).</li>
	<li>Calculate the CL-HB regent Particles Per Cell (PPC) to be added to cells as follows: 
	<img alt="Equation 1" src="/files/ftp_upload/57389/57389eq1.jpg" /> 
	Where the number of cells is the estimated total number of cells at the time of labeling, PPC is the number of Particles Per Cell, and 1 &#215; 108 is the number of particles per mL of the reagent. For example, to label 20,000 cells with a PPC of 30 
	<img alt="Equation 2" src="/files/ftp_upload/57389/57389eq2.jpg" /> 
	NOTE: CL-HB regent works well with most cell types between 10 and 50 PPC. However, 30 PPC worked best for this study.</li>
	<li>Incubate MEFs for 18 h at 37 &#176;C and 5% CO2. 
	NOTE: For the current experiment, visualization of chromosomes in mitotic cells escaping the spindle assembly checkpoint occurred 18 h after cell cycle arrest with nocodazole.</li>
</ol>
5. Visualization of Lagging Chromosomes
<ol>
	<li>Visualize mouse embryonic fibroblasts (MEFs) using a spinning disk confocal microscope system equipped with an environmental chamber and an oil immersion, 63x objective lens.
	<ol>
		<li>Use excitation wavelength of 488 nm and emission wavelength 450 nm for GFP channel image acquisition.</li>
		<li>Use a spinning disk confocal microscope system with the capability of motorized scanning stages, Z-Piezo inserts, stage-top incubation and direct fluorescence recovery after photo bleaching for this technique.</li>
	</ol>
	</li>
	<li>One day before imaging, turn the environmental chamber power ON and warm up the entire chamber at 37 &#176;C overnight.</li>
	<li>Turn power ON for the microscope stand, camera, spinning disk unit, illuminator, argon laser, computer and motorized stage.</li>
	<li>Let the system warm up for 3 min; start the argon laser by turning on ignition key. Switch the toggle switch for argon laser from &#34;standby&#34; to &#34;laser run&#34;.</li>
	<li>Launch the data acquisition and processing software.</li>
	<li>Initiate the CO2 controller for the stage top incubator and set the concentration of CO2 at 5%. This must be done before commencement of imaging.</li>
	<li>Remove the chambered cover glass from the incubator and place on the stage for visualization. Visualize cells via an oil immersion 63x objective lens (NA1.4).</li>
	<li>View through the ocular lenses, focus the image and identify a cell in nuclear envelope breakdown (NEBD) stage.</li>
	<li>Initiate appropriate laser (488 nm argon laser to visualize Histone2B-GFP).</li>
	<li>Open acquisition control window and set exposure time for the GFP channel. Identify a cell that is in NEBD.
	<ol>
		<li>Manually determine the target cell&#39;s top and bottom focal plane, and enter the xyz optical sectioning settings.</li>
		<li>Observe it for 20 min; if the cell does not proceed through cell division, stop image acquisition after 20 min and move on to the next cell that is in NEBD.</li>
	</ol>
	</li>
	<li>Approximately 1 h per cell is needed to image lagging chromosomes during mitosis in PP2A-B56&#947;- cells that escaped from the SAC. To obtain data for a movie, take pictures every 3 min. 
	NOTE: Wild type cells arrest and do not progress past NEBD when treated with nocodazole.</li>
	<li>Save images in zvi file format for further analysis.</li>
</ol>
6. Image Processing and Analysis
Note: Perform three-dimensional image processing and analysis using any available software such as Axiovision version 4.8.2 or Imaris version 8.2. For this study the ImageJ software was used.
<ol>
	<li>Open the image sequence. If the images are already in a stack format, proceed to the next step. If not, combine all relevant images into a stack using &#39;Image &#62; Stacks &#62; Images to Stack&#39; in the menu bar.</li>
	<li>Perform any adjustments as required to brightness/contrast and levels.</li>
	<li>To add a time stamp to the movie:
	<ol>
		<li>Go to &#39;Image &#62; Stacks &#62; Label&#8230;&#39; in the menu bar.</li>
		<li>Select the appropriate format, the starting time value, and time interval between each image.</li>
		<li>Check the &#39;Preview&#39; box and adjust location and format settings. Press &#39;OK&#39; to apply the time stamp.</li>
	</ol>
	</li>
	<li>To add a scale bar to the movie:
	<ol>
		<li>Set the image scaling under &#39;Analyze &#62; Set Scale&#39; in the menu bar.</li>
		<li>In the &#39;Distance in pixels&#39; field, enter a number of pixels where the distance is known, and in the &#39;Known distance&#39; field enter the distance. Set the correct unit of length for the distance, e.g., in &#181;m. Press &#39;OK&#39; to apply the scale to the stack.</li>
		<li>Go to &#39;Analyze &#62; Tools &#62; Scale Bar&#8230;&#39; to add the scale bar. Set the size of the bar as &#39;Width in &#181;m&#39;, and adjust the remaining formatting options as appropriate. Check the &#39;Label all slices&#39; box to add the scale bar to the entire stack.</li>
	</ol>
	</li>
	<li>Preview the movie by clicking on the triangular play button on the bottom left edge of the image window. Adjust the frame rate using &#39;Image &#62; Stacks &#62; Animation &#62; Animation Options&#39; in the menu bar.</li>
	<li>Export the movie file by selecting &#39;File &#62; Save As &#62; AVI&#8230;&#39; and select the frame rate and compression.</li>
	<li>Press &#39;OK&#39; to choose the save location and file name. Press &#39;Save&#39; to save the movie file.</li>
</ol>

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+spindle+assembly+checkpoint.">The spindle assembly checkpoint.</a> Curr Biol. 22, 966-980 (2012).</li><li>Jallepalli, P. V., Lengauer, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chromosome+segregation+and+cancer:+cutting+through+the+mystery.">Chromosome segregation and cancer: cutting through the mystery.</a> Nat Rev Cancer. 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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=Any+Way+You+Slice+It-A+Comparison+of+Confocal+Microscopy+Techniques.">Any Way You Slice It-A Comparison of Confocal Microscopy Techniques.</a> J Biomol Tech. 26 (2), 54-65 (2015).</li><li>Versaevel, M., Braquenier, J. B., Riaz, M., Grevesse, T., Lantoine, J., Gabriele, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Super-resolution+microscopy+reveals+LINC+complex+recruitment+at+nuclear+indentation+sites.">Super-resolution microscopy reveals LINC complex recruitment at nuclear indentation sites.</a> Sci Rep. 8 (4), 7362 (2014).</li><li>Wild, T., Larsen, M. S., Narita, T., Schou, J., Nilsson, J., Choudhary, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Spindle+Assembly+Checkpoint+Is+Not+Essential+for+Viability+of+Human+Cells+with+Genetically+Lowered+APC/C+Activity.">The Spindle Assembly Checkpoint Is Not Essential for Viability of Human Cells with Genetically Lowered APC/C Activity.</a> Cell Rep. 1 (8), 1829-1840 (2016).</li><li>Iuso, D., 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=Exogenous+Expression+of+Human+Protamine+1+(hPrm1)+Remodels+Fibroblast+Nuclei+into+Spermatid-like+Structures.">Exogenous Expression of Human Protamine 1 (hPrm1) Remodels Fibroblast Nuclei into Spermatid-like Structures.</a> Cell Rep. 1 (9), 1765-1771 (2015).</li><li>Ratcliffe, E., Glen, K. E., Naing, M. W., Williams, D. 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The authors declare that they have no competing financial interests.

Cell cycle control checkpoints that ensure accurate chromosome segregation prevent aneuploidy and cell transformation 1,2,3. In the present study, we found that inactivation of PP2A-B56&#947; resulted in a weakened spindle assembly checkpoint. Live cell imaging allowed us to observe chromosomal mis-segregation during mitosis in PP2A-B56&#947; MEFs treated with nocodazole 9.
This protocol utilizes Histone2B-GFP label generated by BacMam 2.0 technology to label chromosomes for live cell imaging. One of the critical steps in labeling is calculating right amount of particles per cells (PPC) to achieve enough labeling. It was found through titration that 10-50 PPC works best. This reagent could be toxic to the cells; hence it is necessary to titrate PPC used to achieve balance between labelling and toxicity to the labelled cells. Also, this is a transient transfection and can be detected only up to 5 days, hence, it may not be appropriate for studies that would warrant longer observation periods.
Traditional tagging of the protein or DNA can require complicated protocols, but here we use a reagent which requires only an overnight incubation with desired cells. In addition, live cell imaging was performed using spinning disc confocal microscopy which unlike confocal laser scanning microscopy is faster and reduces phototoxicity and bleaching effects in the living cells 11. This protocol allows easy, convenient method to label mitotic cells and visualize chromosomal segregation as compared to the other techniques such as FISH, immunohistochemistry and flow cytometry. This protocol also allows the capture of lagging chromosomes images throughout mitosis providing additional information to that obtained using traditional immunocytochemistry, FISH or flow cytometry, thereby making it possible to capture the phenotypes such as lagging chromosome in both space and time. This protocol allows convenient, fast and easy labeling of chromosomes in wide variety of cell lines, primary cells, stem cells, neurons and immortalized T-cells 12,13,14.
Various types of cell therapy products are being developed with the hopes to treat varied disease conditions 15. One of the challenges faced in the development of these cell therapies involves safety concerns due to possibility of genetic instability and tumorigenesis. This protocol describes a convenient and easy method to label chromosomes in variety of cell types. In the future it could be employed to develop an assay to study and measure genetic instability and quality of the cell therapy products.

In the following protocol, we describe a convenient method to visualize the chromosomal segregation and mitotic progression during cell cycle in mouse embryonic fibroblasts using Histone 2B-GFP, BacMam 2.0 labeling and live cell imaging.
Cell cycle control checkpoints monitor chromosome segregation and play an important role in the maintenance of the genetic integrity of the cell 1,2,3. Accumulation of mis-segregated chromosomes can lead to aneuploidy, which is a hallmark of most solid tumors 4. Hence, detection of lagging chromosomes can be used as a method to study chromosomal instability.
Fluorescently labeled proteins can be used to visualize live chromosome segregation and chromosomal lagging but the generation of mCherry-tagged or H2B-GFP tagged protein requires substantial knowledge of gene delivery and molecular biology 5. Here we describe the use of CellLight Histone2B-GFP BacMam 2.0 reagent, hereafter called CL-HB regent, for the sake of simplicity. This reagent can be used immediately and thus eliminates concerns about vector quality and integrity. In addition, this reagent does not require the use of potentially harmful treatments or lipids and dye-loading chemicals. Unlike conventional fluorescent labels, the CL-HB regent stains independently of function (i.e., membrane potential). The CL-HB regent can be simply added to the cells and incubated overnight for protein expression. The CL-HB regent does not replicate in mammalian cells and can be used in biosafety level (BSL) 1 laboratory settings. Also, this transient transfection can be detected after overnight incubation for up to 5 days, enough time to carry out most dynamic cellular analyses.
Alternatively, chromosomal abnormalities could be studied by various techniques such as flow cytometry, immunohistochemistry or fluorescence in situ hybridization (FISH) 6. Flow cytometry can be used to study aneuploidy, which can be measured based on DNA content and the phase of cells in the cell cycle. Although flow cytometry can be used to measure aneuploidy, it does not provide information on chromosomal mis-segregation. FISH and immunohistochemistry techniques use fluorescent probes to bind to DNA or chromosomes. While these techniques provide a snapshot of the status of a population of cells, they do not allow live cell imaging thereby missing any information obtained through spatiotemporal visualization of cytokinesis in specific cells followed over a period of time.
This protocol was used to study lagging chromosomes or chromosomal mis-segregation in nocodazole treated mouse embryonic fibroblasts (MEFs) isolated from PP2A-B56&#947;- mice. In addition to above application, this protocol provides a simple tool to label and visualize chromosomal segregation in various cell types which can be used to study cell cycle regulation or chromosomal instability in tumor cells. In addition, it can also be used to study chromosomal instability caused by various drug treatments or to study the effects of gene knock out resulting in chromosomal mis-segregation.

<table><tbody><tr><td>DMEM/F12</td><td>Gibco</td><td>11320082</td><td></td></tr><tr><td>L-Glutamine</td><td>Gibco</td><td>25030081</td><td></td></tr><tr><td>MEM Non-Essential amino acids solution (100X)</td><td>Gibco</td><td>11140050</td><td></td></tr><tr><td>Penicillin-Streptomycin (10,000 U/mL)</td><td>Gibco</td><td>15140122</td><td></td></tr><tr><td>Dulbecco&amp;rsquo;s Phosphate Buffered saline</td><td>Gibco</td><td>14190144</td><td></td></tr><tr><td>Trypsin-EDTA</td><td>Gibco</td><td>25300054</td><td></td></tr><tr><td>Fetal bovine serum</td><td>Atlanta Biologicals</td><td>S11150</td><td></td></tr><tr><td>DMSO</td><td>EMD Millipore</td><td>MX 1458-6</td><td></td></tr><tr><td>Cryogenic vial storage boxes</td><td>Fisherbrand</td><td>10-500-28</td><td></td></tr><tr><td>Cryogenic vials</td><td>Corning/costar</td><td>431416</td><td></td></tr><tr><td>T75 flasks</td><td>Cellstar</td><td>658175</td><td></td></tr><tr><td>2 well chambered cover glass</td><td>Nunc</td><td>155380PK</td><td></td></tr><tr><td>Cellometer Vision Cell Profiler</td><td>Nexcelom Bioscience LLC</td><td>Cellometer Vision Trio</td><td></td></tr><tr><td>Nocodazole</td><td>Sigma</td><td>M1404</td><td></td></tr><tr><td>CellLight Histone 2B-GFP, BacMam 2.0</td><td>Thermo Fisher Scientific Inc.</td><td>C10594</td><td></td></tr><tr><td>Zeiss Cell Observer Spinning Disk Confocal Microscope system</td><td>Carl Zeiss Microscopy</td><td>Zeiss Cell Observer SD</td><td></td></tr><tr><td>Water Bath</td><td>Thermoscientific</td><td>280 series</td><td></td></tr><tr><td>Incubator</td><td>Sanyo commercial solutions</td><td>MCO-18AIC (UV)</td><td></td></tr><tr><td>Class II Biological safety cabinet</td><td>The Baker Company</td><td>SterilGard</td><td></td></tr><tr><td>Axiovision software</td><td>Zeiss</td><td>Ver.4.8.2</td><td></td></tr><tr><td>ImageJ software</td><td>National Institute of Health</td><td>Ver. 1.51r</td><td></td></tr></tbody></table>

live cell imaging of chromosome segregation during mitosis

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled by multiple mechanisms that include the Spindle Assembly Checkpoint (SAC). The SAC is part of a complex feedback system that is responsible for prevention of a cell progress through mitosis unless all chromosomal kinetochores have attached to spindle microtubules. Chromosomal lagging and abnormal chromosome segregation is an indicator of dysfunctional cell cycle control checkpoints and can be used to measure the genomic stability of dividing cells. Deregulation of the SAC can result in the transformation of a normal cell into a malignant cell through the accumulation of errors during chromosomal segregation. Implementation of the SAC and the formation of the kinetochore complex are tightly regulated by interactions between kinases and phosphatase such as Protein Phosphatase 2A (PP2A). This protocol describes live cell imaging of lagging chromosomes in mouse embryonic fibroblasts isolated from mice that had a knockout of the PP2A-B56&#947; regulatory subunit. This method overcomes the shortcomings of other cell cycle control imaging techniques such as flow cytometry or immunocytochemistry that only provide a snapshot of a cell cytokinesis status, instead of a dynamic spatiotemporal visualization of chromosomes during mitosis.

MEFs from wild type and PP2A-B56&#947;- mice were seeded in a 2-well chamber cover glass and allowed to attach. On day 2, MEFs were synchronized using 0.1% FBS for 24 h. On day 3, MEFs in media with 200 ng/mL nocodazole and 30 PPC CL-HB regentwere incubated for 18 h at 37&#176;C and 5% CO2. On day 4, the cells were imaged using a spinning disk confocal microscope system (Figure 1). Live cell imaging was used to visualize the fate of individual cells as they progressed from NEBD through mitosis (Figure 2). Nocodazole is known to arrest the wild type cells at G2M phase by interfering with microtubule formation. It was observed that post nocodazole treatment wild type cells were arrested at metaphase upper panel (Figure 2). We observed that lack of PP2A-B56&#947; weakened the cell cycle control, which allowed PP2A-B56&#947;- MEFs to overcome cell cycle arrest induced by nocodazole 9. In addition, abnormal chromosomal segregation was detected in 62% (13 out of 21) of mitotic PP2A-B56&#947;- MEFs treated with nocodazole (shown by yellow arrows in lower panel in Figure 2). Five of the PP2A-B56&#947;- MEFs arrested at NEBD and 3 went through mitosis without observable defects. In contrast, all 21 wild type cells that were imaged did not proceed through mitosis and were found to be arrested by metaphase (upper panel Figure 2).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57389/57389fig1.jpg" /> 
Figure 1. Basic flowchart for live cell imaging. MEFs from wild type and PP2A-B56&#947;- mice were seeded in a 2-well chambered cover glass overnight. Day 2, cells were synchronized using 0.1% FBS for 24 h. Day 3, cells were treated with 200 ng/mL nocodazole in growth media and 30 PPC Histone 2B-GFP, BacMam 2.0 reagent for 18 h. Day 4 the cells were imaged using spinning disk confocal microscope system. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57389/57389fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57389/57389fig2.jpg" /> 
Figure 2. Chromosomal abnormalities are detected in PP2A-B56&#947;- MEFs. Time lapse microscopy live images captured at various time points in mitosis (0-51 min) of nocodazole treated wild type and PP2A-B56&#947;- MEFs. The wild type cells were found to be arrested at metaphase (upper panel), whereas PP2A-B56&#947;- MEFs were able to go through mitosis (lower panel). Chromosomal lagging or mis-segregation can be seen in PP2A-B56&#947;- MEFs shown by yellow arrows in the lower panel. Scale bar = 5 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57389/57389fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Live Cell Imaging of Chromosome Segregation During Mitosis at JoVE.com

Live Cell Imaging of Chromosome Segregation During Mitosis

This protocol describes an easy and convenient method to label and visualize live chromosomes in mitotic cells using Histone2B-GFP BacMam 2.0 label and a spinning disc confocal microscopy system.

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled ...

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9.2K Views.  US Food and Drug Administration. This protocol describes an easy and convenient method to label and visualize live chromosomes in mitotic cells using Histone2B-GFP BacMam 2.0 label and a spinning disc confocal microscopy system.

Video: Live Cell Imaging of Chromosome Segregation During Mitosis

Long-term Live-cell Imaging to Assess Cell Fate in Response to Paclitaxel

Live-cell imaging is a powerful technique that can be used to directly visualize biological phenomena in single cells over extended periods of time. Over the past decade, new and innovative technologies have greatly enhanced the practicality of live-cell imaging. Cells can now be kept in focus and continuously imaged over several days while maintained under 37 &#176;C and 5% CO2 cell culture conditions. Moreover, multiple fields of view representing different experimental conditions can be acquired simultaneously, thus providing high-throughput experimental data. Live-cell imaging provides a significant advantage over fixed-cell imaging by allowing for the direct visualization and temporal quantitation of dynamic cellular events. Live-cell imaging can also identify variation in the behavior of single cells that would otherwise have been missed using population-based assays. Here, we describe live-cell imaging protocols to assess cell fate decisions following treatment with the anti-mitotic drug paclitaxel. We demonstrate methods to visualize whether mitotically arrested cells die directly from mitosis or slip back into interphase. We also describe how the fluorescent ubiquitination-based cell cycle indicator (FUCCI) system can be used to assess the fraction of interphase cells born from mitotic slippage that are capable of re-entering the cell cycle. Finally, we describe a live-cell imaging method to identify nuclear envelope rupture events.

Live-cell imaging provides a wealth of information on single cells or whole populations that is unattainable by fixed cell imaging alone. Here, live-cell imaging protocols to assess cell fate decisions following treatment with the anti-mitotic drug paclitaxel are described.

Live-cell imaging is a powerful technique that can be used to directly visualize biological phenomena in single cells over extended periods of time. Over ...

Cancer Research

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a rapid and high-throughput manner. S2 cells have proven amenable to both fixed- and live-cell imaging applications. Notably, live-cell imaging can yield valuable information about how loss or knockdown of a gene can affect the kinetics and dynamics of key events during cell division, including mitotic spindle assembly, chromosome congression, and segregation, as well as overall cell cycle timing. Here we utilize S2 cells stably transfected with fluorescently tagged mCherry:&#945;-tubulin to mark the mitotic spindle and GFP:CENP-A (referred to as &#8216;CID&#8217; gene in Drosophila) to mark the centromere to analyze the effects of key mitotic genes on the timing of cell divisions, from prophase (specifically at Nuclear Envelope Breakdown; NEBD) to the onset of anaphase. This imaging protocol also allows for the visualization of the spindle microtubule and chromosome dynamics throughout mitosis. Herein, we aim to provide a simple yet comprehensive protocol that will allow readers to easily adapt S2 cells for live imaging experiments. Results obtained from such experiments should expand our understanding of genes involved in the cell division by defining their role in several simultaneous and dynamic events. Observations made in this cell culture system can be validated and further investigated in vivo using the impressive toolkit of genetic approaches in flies.

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Cell divisions can be visualized in real time using fluorescently tagged proteins and time-lapse microscopy. Using the protocol presented here, users can analyze cell division timing dynamics, mitotic spindle assembly, and chromosome congression and segregation. Defects in these events following RNA interference (RNAi)-mediated gene knockdown can be assessed and quantified.

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a ...

Genetics

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Neuroscience

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 ...

Biology

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 ...

Combining Mitotic Cell Synchronization and High Resolution Confocal Microscopy to Study the Role of Multifunctional Cell Cycle Proteins During Mitosis

Study of the various regulatory events of the cell cycle in a phase-dependent manner provides a clear understanding about cell growth and division. The synchronization of cell populations at specific stages of the cell cycle has been found to be very useful in such experimental endeavors. Synchronization of cells by treatment with chemicals that are relatively less toxic can be advantageous over the use of pharmacological inhibitory drugs for the study of consequent cell cycle events and to obtain specific enrichment of selected mitotic stages. Here, we describe the protocol for synchronizing human cells at different stages of the cell cycle, including both in S phase and M phase with a double thymidine block and release procedure for studying the functionality of mitotic proteins in chromosome alignment and segregation. This protocol has been extremely useful for studying the mitotic roles of multifunctional proteins which possess established interphase functions. In our case, the mitotic role of Cdt1, a protein critical for replication origin licensing in G1 phase, can be studied effectively only when G2/M-specific Cdt1 can be depleted. We describe the detailed protocol for depletion of G2/M-specific Cdt1 using double thymidine synchronization. We also explain the protocol of cell fixation, and live cell imaging using high resolution confocal microscopy after thymidine release. The method is also useful for analyzing the function of mitotic proteins under both physiological and perturbed conditions such as for Hec1, a component of the Ndc80 complex, as it enables one to obtain large sample sizes of mitotic cells for fixed and live cell analysis as we show here.&#160;&#160;

We present a protocol for double thymidine synchronization of HeLa cells followed by analysis using high resolution confocal microscopy. This method is key to obtaining large number of cells that proceed synchronously from S phase to mitosis, enabling studies on mitotic roles of multifunctional proteins which also possess interphase functions.

Study of the various regulatory events of the cell cycle in a phase-dependent manner provides a clear understanding about cell growth and division. The ...

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, ...

Super-Resolution Live Cell Imaging of Subcellular Structures

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally been restricted to be used only on fixed samples or live cells outside of tissue labeled with strong fluorescent signal. Current super-resolution live cell imaging techniques require the use of special fluorescence probes, high illumination, multiple image acquisitions with post-acquisition processing, or often a combination of these processes. These prerequisites significantly limit the biological samples and contexts that this technique can be applied to.
Here we describe a method to perform super-resolution (~140 nm XY-resolution) time-lapse fluorescence live cell imaging in situ. This technique is also compatible with low fluorescent intensity, for example, EGFP or mCherry endogenously tagged at lowly expressed genes. As a proof-of-principle, we have used this method to visualize multiple subcellular structures in the Drosophila testis. During tissue preparation, both the cellular structure and tissue morphology are maintained within the dissected testis. Here, we use this technique to image microtubule dynamics, the interactions between microtubules and the nuclear membrane, as well as the attachment of microtubules to centromeres.
This technique requires special procedures in sample preparation, sample mounting and immobilizing of specimens. Additionally, the specimens must be maintained for several hours after dissection without compromising cellular function and activity. While we have optimized the conditions for live super-resolution imaging specifically in Drosophila male germline stem cells (GSCs) and progenitor germ cells in dissected testis tissue, this technique is broadly applicable to a variety of different cell types. The ability to observe cells under their physiological conditions without sacrificing either spatial or temporal resolution will serve as an invaluable tool to researchers seeking to address crucial questions in cell biology.

Presented here is a protocol for super-resolution live-cell imaging in intact tissue. We have standardized the conditions for imaging a highly sensitive adult stem cell population in its native tissue environment. This technique involves balancing temporal and spatial resolution to allow for the direct observation of biological phenomena in live tissue.

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally ...

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 ...

Live Cell Imaging of Chromosome Segregation During Mitosis

Summary

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Chapters in this video

Long-term Live-cell Imaging to Assess Cell Fate in Response to Paclitaxel

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Time-lapse Imaging of Mitosis After siRNA Transfection

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

Combining Mitotic Cell Synchronization and High Resolution Confocal Microscopy to Study the Role of Multifunctional Cell Cycle Proteins During Mitosis

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

Super-Resolution Live Cell Imaging of Subcellular Structures

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