We are grateful to Dr. Kozo Tanaka of Tohoku University, Japan for sharing HeLa cells stably expressing mCherry-Histone H2B and GFP-&#945;-tubulin. This work was supported by an NCI grant to DV (R00CA178188) and by start-up funds from Northwestern University.

1. Double Thymidine Block and Release: Reagent Preparations
<ol>
	<li>Make 500 mL Dulbecco&#39;s modified Eagle medium (DMEM) medium supplemented with 10% FBS, penicillin, and streptomycin.</li>
	<li>Make 100 mM stock of thymidine in sterile water and store in aliquots at -80 &#176;C.</li>
</ol>
2. Protocol for Fixed Cell Imaging of Mitotic Progression (Figure 1A)
<ol>
	<li>On day 1, seed ~2 x 105 HeLa cells into the wells of a 6-well plate with a cover slip (sterilized with 70% ethanol and UV irradiation) and 2 mL of DMEM medium. Grow the cells in a humidified incubator for 24 h at 37 &#176;C and 5% CO2.</li>
	<li>1st thymidine block: On day 2, thoroughly mix the required volume of thymidine in fresh DMEM media (100 mM stock, 2 mM final concentration).</li>
	<li>Add 2 mL of thymidine-containing media to the cells in each well of the 6-well plate and incubate for 18 h.</li>
	<li>On day 3, aspirate the medium and wash the cells twice with 2 mL of 1x PBS and once with fresh prewarmed DMEM; grow cells in fresh prewarmed DMEM medium for 9 h to release cells from block.</li>
	<li>2nd thymidine block: Again add 2 mL of thymidine-containing media (2 mM final concentration) to the cells in each well of the 6-well plate.</li>
	<li>Incubate for another 18 h.</li>
	<li>On day 4, aspirate the medium and wash the cells twice with 2 mL of 1x PBS and once with fresh prewarmed DMEM.</li>
	<li>Dilute siRNAs (control and Cdt1) and transfection reagent in serum free growth medium separately for 10 min. Mix them together and incubate for 20 min at RT. The final concentration of siRNA used in each transfection was 100 nM. Add the reaction mixture to the cells that have been washed out of Thymidine.</li>
	<li>Incubate cells at 37 &#176;C for 9-10 h to release from blocking and fix the cells on the cover slips with 4% PFA for 20 min at RT. 
	Caution: PFA is toxic, wear appropriate protection.</li>
	<li>Immunostain cells, after permeabilizing with 0.5% (v/v) detergent for 10 min at RT and washing the cells twice with 1x PBS for 5 min each.
	<ol>
		<li>Block cells with 1% BSA (w/v) in 1x PBS for 1 h at RT followed by treating cells with 50 &#181;L of primary antibodies (mouse anti-&#945;-tubulin antibody diluted at 1:1,000, rabbit anti-Zwint1 antibody diluted at 1:400, and mouse anti-phospho-&#611; H2AX diluted at 1:300) in 1% (w/v) BSA in 1x PBS for 1 h at 37 oC.</li>
		<li>After washing out the cells with 1x PBS thrice, treat cells with 50 &#181;L of secondary antibodies for each cover glass (Alexa 488 and Rhodamine Red at 1:250 dilutions in 1% (w/v) BSA in 1x PBS) for 1 h at RT.</li>
		<li>After washing the cells with 1x PBS twice for 5 min each, treat cells with DAPI (0.1&#181;g/mL in 1x PBS) for 5 min at RT.</li>
		<li>After washing the cells with 1x PBS twice for 5 min each, place the cover slip facing cells to the appropriate mounting media on a clear microscopic slide.</li>
	</ol>
	</li>
	<li>Image the cells for the immunostained proteins with 60X or 100X 1.4 NA Plan-Apochromatic DIC oil immersion objective mounted on an inverted high resolution confocal microscope equipped with an appropriate camera as necessary for the quality of images required.</li>
	<li>Acquire images at room temperature as z-stacks of 0.2 &#181;m thickness using software appropriate for the microscope.</li>
</ol>
3. Protocol for Live Cell Imaging of Mitotic Progression (Figure 3A)
<ol>
	<li>On day 1, seed approximately 0.5-1 x 105 HeLa cells stably expressing mCherry-H2B and GFP-&#945;-tubulin (slightly variable based on cell type and the proteins they express) into 35 mm glass bottom dishes with 1.5 mL of DMEM medium and grow them in a humidified incubator for 24 h at 37 oC and 5% CO2.</li>
	<li>1st thymidine block: On day 2, add 1.5 mL of thymidine-containing media to the cells in the dish (100 mM thymidine, 2 mM final concentration,).</li>
	<li>Incubate for 18 h.</li>
	<li>On day 3, aspirate the medium containing thymidine and wash the cells thrice, twice with 2 mL of 1x PBS and once with fresh prewarmed DMEM.</li>
	<li>Dilute siRNAs (control and Hec1) and transfection reagent with serum free growth medium separately for 10 min. Mix them together and incubate for 20 min at RT. The final concentration of siRNA used in each transfection was 100 nM. Add the reaction mixture to the cells that have been washed out of Thymidine.</li>
	<li>Grow cells in 1.5 mL of fresh prewarmed DMEM medium for 8 h to release cells from block.</li>
	<li>2nd thymidine block: Add 1.5 mL of thymidine-containing media (2 mM final concentration) to the cells in the dish.</li>
	<li>Incubate for another 18 h.</li>
	<li>On day 4, aspirate the medium and wash the cells thrice, twice with 2 mL of 1x PBS and once with fresh prewarmed DMEM. Then grow cells in fresh prewarmed Leibovitz&#39;s (L-15) medium supplemented with 10% FBS and 20 mM HEPES at pH 7.0 for 8 h to release cells from the block.</li>
	<li>Place the dish in the temperature control chamber set in the high resolution confocal microscope which has already switched on at least 30 min before beginning the imaging to stabilize the on-stage and experimental temperatures.</li>
	<li>Focus in bright field with 60x objective until cells are visible, then manually sift the on-stage to the region of choice.</li>
	<li>Set up the light and laser power/exposure, image acquisition parameters, and duration of the experiment using the microscope image acquisition software of choice. Use filters for GFP (488 excitation; 385 nm emission) and mCherry (561 nm excitation and 385 nm emission) to acquire images.</li>
	<li>Perform the time-lapse experiment by separately acquiring pulsed transmitted light and fluorescence images every 10 min for a period up to 16 h.</li>
	<li>Acquire images as twelve 1.0 &#181;m-separated z-planes at 9 h after release from the double thymidine block using software appropriate to the microscope.</li>
	<li>Analyze the images tracking individual cells for mitotic progression and finally assemble the corresponding movie with the source software Fiji/ImageJ.</li>
</ol>

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The authors declare that they have no competing financial interest.

The most critical advantage of double thymidine synchronization is that it provides an increased sample size of mitotic cells in a short time window with many of these cells entering mitosis in unison, thus also enabling analyses of chromosome alignment, bipolar spindle formation, and chromosome segregation with much higher efficiency.
Many regulatory protein complexes and signaling pathways are devoted to ensuring normal progression through mitosis and deregulation of this process may led to tumorigenesis. Live cell imaging of HeLa cells stably expressing mCherry-H2B and GFP-tubulin and released from the double thymidine block shows that most control cells segregate their chromosomes at anaphase within approximately 60 min after NEB (Figure 3B). On the other hand, live cell imaging of HeLa cells perturbed with Hec1 function shows that most cells stay in the mitotic phase for a long period with misaligned chromosomes followed by anaphase onset or apoptosis (Figure 3C).
Excess thymidine inhibits DNA synthesis during the S phase, thus blocking cells at G1/S phase11. Although double thymidine block synchronization is the most commonly used method in the study of the cell cycle and mitotic progression, one needs to be critical of some of the limitations of this method. Importantly, it needs to be ensured that the chemical has been washed out thoroughly (for 7-8 h) so that the effect is completely reversible and cells can proceed normally through the rest of the cell cycle. Cells may otherwise not enter the second cycle causing improper progression though mitosis. The proportion of synchronized cells may not be sufficiently high with a single thymidine block and a double thymidine block may be necessary to increase the number of synchronized cells, especially if one is interested in studying a specific stage of mitosis. The duration of the thymidine treatment must be optimized due to different sensitivity among cell types depending on the generation time. For example, hamster cells have 16 h of generation time and are treated with thymidine for 11 h for optimal synchronization17. In this study, we treated HeLa cells for 16-18 h with thymidine. Moreover, the excess thymidine not only prevents DNA synthesis but also may kill important fractions of the cells11,12,18 and close attention needs to be paid to monitor this loss. The thymidine stock solution used in this study should be prepared in sterile distilled water and should be mixed thoroughly in the cell culture media to avoid its precipitation after adding it to media of siRNA transfected cells and also to ensure its homogenous distribution around the cultured cells.
For studying the function of a target mitotic protein during mitotic progression, the cells would ideally be treated with siRNA to knockdown the levels of the protein of interest either during the 1st thymidine block or during the release from the 1st thymidine block, depending upon the time required to obtain efficient protein knockdown19,20,21. On the other hand, for studying the role of multifunctional proteins such as Cdt1 during mitosis, it is necessary to treat the cells with siCdt1 during the release from the 2nd thymidine block to obtain G2/M specific depletion. Cdt1 is a critical player in the licensing of DNA replication origins15. To understand its specialized role during mitosis without interfering with its role in licensing, Cdt1 needs to be depleted specifically at G2/M phase, which can be efficiently achieved by double thymidine synchronization, thus avoiding the need to resort to other mitosis-specific perturbation approaches that are difficult to execute. Depletion of Cdt1 in asynchronous cultures induce DNA damage response due to the interruption of the licensing of DNA replication origins, which would in turn have muddled the analysis of a possible function for this protein during mitosis. Thus, the double thymidine synchronization technique offers a unique experimental approach to generate cells that underwent a normal G1 and S phase but lacked Cdt1 function during G2 and M phase. The special significance of our synchronization protocol lies in its inhibition of multifunctional proteins at the specific phase (G2/M) to study their novel functions during mitosis. Thus, this method will be useful not only to study the role of proteins that have multiple functions during different cell cycle stages but also of that of any mitotic protein in the process of chromosome alignment, kMT attachments, and chromosome segregation.
For live cell imaging, after release from the double thymidine block, cells should be incubated in pre-warmed Leibovitz&#39;s (L-15) medium supplemented with 10-20% FBS until the beginning of imaging. The cells enter mitosis at variable time points after release from thymidine block depending on the cell type. The start time of image capturing or fixation needs to be optimized depending on the cell type. For live cell imaging, the utilization of conventional confocal microscopy may cause photobleaching during long-term imaging. On the other hand, the high resolution confocal microcopy gives a clear and high-resolution image with the least extent of photobleaching effects.
The critical steps in this protocol are the duration of thymidine treatment, proper washout of the thymidine, the timing of siRNA transfection, and the start timing of imaging.

In the cell cycle, cells undergo a series of highly regulated and temporally controlled events for the accurate duplication of their genome and proliferation. In mammals, the cell cycle consists of interphase and M-phase. In interphase, which consists of three stages- G1, S, and G2, the cell duplicates its genome and undergoes growth that is necessary for normal cell cycle progression1,2. In the M-phase, which consists of mitosis (prophase, prometaphase, metaphase, anaphase, and telophase) and cytokinesis, a parental cell produces two genetically identical daughter cells. In mitosis, sister chromatids of duplicated genome are condensed (prophase) and are captured at their kinetochores by microtubules of the assembled mitotic spindle (prometaphase), that drives their alignment at the metaphase plate (metaphase) followed by their equal segregation when sister chromatids are split toward and transported to opposite spindle poles (anaphase). The two daughter cells are physically separated by the activity of an actin-based contractile ring (telophase and cytokinesis). The kinetochore is a specialized proteinaceous structure which assembles at the centromeric region of chromatids and serve as attachment sites for spindle microtubules. Its main function is to drive chromosome capture, alignment, and aid in correcting improper spindle microtubule attachment, while mediating the spindle assembly checkpoint to maintain the fidelity of chromosome segregation3,4.
The technique of cell synchronization serves as an ideal tool for understanding the molecular and structural events involved in cell cycle progression. This approach has been used to enrich cell populations at specific phases for various types of analyses, including profiling of gene expression, analyses of cellular biochemical processes, and detection of subcellular localization of proteins. Synchronized mammalian cells can be used not only for the study of individual gene products, but also for approaches involving analysis of whole genomes including microarray analysis of gene expression5, miRNA expression patterns6, translational regulation7, and proteomic analysis of protein modifications8. Synchronization can also be used to study the effects of gene expression or protein knock-down or knock-out, or of chemicals on cell cycle progression.
Cells can be synchronized at the different stages of the cell cycle. Both physical and chemical methods are widely used for cell synchronization. The most important criteria for cell synchronization are that synchronization should be noncytotoxic and reversible. Because of the potential adverse cellular consequences of synchronizing cells by pharmacological agents, chemical-dependent methods can be advantageous for studying key cell cycle events. For example, hydroxyurea, amphidicolin, mimosine, and lovastatin, can be used for cell synchronization at G1/S phase but, because of their effect on the biochemical pathways they inhibit, they activate cell cycle checkpoint mechanisms and kill an important fraction of the cells9,10. On the other hand, feedback inhibition of DNA replication by adding thymidine to the growth media, known as &#34;thymidine block&#34;, can arrest the cell cycle at certain points11,12,13. Cells can also be synchronized at G2/M phase by treating with nocodazole and RO-33069,14.Nocodazole, which prevents microtubule assembly, has a relatively high cytotoxicity. Moreover, nocodazole-arrested cells can return to interphase precociously by mitotic slippage. Double thymidine block arrest cells at G1/S phase and after release from the block, cells are found to proceed synchronously through G2 and into mitosis. The normal progression of the cell cycle for cells released from thymidine block can be observed under high resolution confocal microscopy by either cell fixation or live imaging. The effect of perturbation of mitotic proteins can be studied specifically when cells enter and proceed through mitosis after release from double thymidine block. Cdt1, a multifunctional protein, is involved in DNA replication origin licensing in the G1 phase and is also required for kinetochore microtubule attachments during mitosis15. To study the function of Cdt1 during mitosis, one needs to adopt a method that avoids the effect of its depletion on replication licensing during G1 phase, while at the same time effecting its depletion specifically during the G2/M phase only. Here, we present detailed protocols based on the double thymidine block to study the mitotic role of proteins performing multiple functions during different stages of the cell cycle by both fixed and live-cell imaging.

<table><tbody><tr><td>DMEM (1x)</td><td>Life Technologies</td><td>11965-092</td><td>Store at 4 &amp;deg;C</td></tr><tr><td>DPBS (1x)</td><td>Life Technologies</td><td>14190-144</td><td>Store at 4 &amp;deg;C</td></tr><tr><td>Leibovitz&amp;rsquo;s (1x) L-15 medium</td><td>Life Technologies</td><td>21083-027</td><td>Store at 4 &amp;deg;C</td></tr><tr><td>Serum reduced medium (Opti-MEM)</td><td>Life Technologies</td><td>319-85-070</td><td>Store at 4 &amp;deg;C</td></tr><tr><td>Penicillin and streptomycin</td><td>Life Technologies</td><td>15070-063 (Pen Strep)</td><td>1:1,000 dilution</td></tr><tr><td>Dharmafect2</td><td>GE Dharmacon</td><td>T-2002-02</td><td>Store at 4 &amp;deg;C</td></tr><tr><td>Thymidine</td><td>MP Biomedicals LLC</td><td>103056</td><td>Dissolved in sterile distiled water</td></tr><tr><td>Cdt1 siRNA</td><td>Life Technologies</td><td></td><td>Ref 10</td></tr><tr><td>Hec1 siRNA</td><td>Life Technologies</td><td></td><td>Ref 13</td></tr><tr><td>HeLa cells expressing GFP-H2B</td><td></td><td></td><td></td></tr><tr><td>HeLa cells expressing GFP-&amp;alpha;-tubulin and mCherry H2B</td><td>Generous gift from Dr. Kozo Tanaka of Tohoku university, Japan</td><td></td><td></td></tr><tr><td>Formaldehyde solution</td><td>Sigma-Aldrich Corporation</td><td>F8775</td><td>Toxic, needs caution</td></tr><tr><td>DAPI</td><td>Sigma-Aldrich Corporation</td><td>D9542</td><td>Toxic, needs caution</td></tr><tr><td>Mouse anti-&amp;alpha;-tubulin</td><td>Santa Cruz Biotechnology</td><td>Sc32293</td><td>1:1,000 dilution</td></tr><tr><td>Rabbit ant-Zwint1</td><td>Bethyl</td><td>A300-781A</td><td>1:400 dilution</td></tr><tr><td>Mouse anti-phospho-&amp;gamma;H2AX (Ser139)</td><td>Upstate Biotechnology</td><td>05-626, clone JBW301</td><td>1:300 dilution</td></tr><tr><td>Alexa 488</td><td>Jackson ImmunoResearch</td><td></td><td>1:250 dilution</td></tr><tr><td>Rodamine Red-X</td><td>Jackson ImmunoResearch</td><td></td><td>1:250 dilution</td></tr><tr><td>BioLite 6 well multidish</td><td>Thermo Fisher Scientific</td><td>130184</td><td></td></tr><tr><td>35 mm Glass bottom dish</td><td>MatTek Corporation</td><td>P35GCOL-1.5-14-C</td><td></td></tr><tr><td>Nikon Eclipse TiE inverted microscope</td><td>Nikon Instruments</td><td></td><td></td></tr><tr><td>Spinning disc for confocal</td><td>Yokagawa</td><td>CSU-X1</td><td></td></tr><tr><td>Ultra 888 EM-CCD Camera</td><td>Andor</td><td>iXon Ultra EMCCD</td><td></td></tr><tr><td>4 wave length laser</td><td>Agilent Technologies</td><td></td><td></td></tr><tr><td>Incubation System for Microscopes</td><td>Tokai Hit</td><td>TIZB</td><td></td></tr><tr><td>NIS-elements software</td><td>Nikon Instruments</td><td></td><td></td></tr></tbody></table>

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;

Study of mitotic progression and microtubule stability in cells fixed after release from double thymidine block 
Cdt1 is involved in licensing of DNA replication origins in the G1 phase. It is degraded during S phase but re-accumulates in G2/M phase. To study its role in mitosis, the endogenous Cdt1 needs to be depleted specifically at the G/M phase using the most suitable cell synchronization technique, the double thymidine block15. Cells were transfected with siRNAs immediately after release from the 2nd thymidine block, when cells are in the S phase and by which time the licensing of replication origins in G1, which also requires Cdt1 function, has long been completed. The depletion of Cdt1 after 9 h of siCdt1 transfection and double thymidine block release was validated by Western blotting (Figure 1B). Fixation of cells 9 h after release from the double thymidine block shows that most cells are at the metaphase stage in both control and Cdt1 siRNA transfected cells. But fixation of cells 10 h after release from the double thymidine block shows that most Cdt1 siRNA-treated cells are still arrested in late prometaphase stage while control cells enter anaphase and normally segregate their chromosomes as expected (Figure 1C). To understand the effect of Cdt1 depletion on kinetochore-microtubule (kMT) stability, cells were fixed at 9 h after release from double thymidine block followed by cold treatment, which only retains stable kinetochore-microtubules (kMTs). The kMTs of Cdt1-depleted cells are relatively less robust after cold treatment compared to that of control cells (Figure 1D). Thus, using this approach, the progression of normal mitotic cells after the perturbation of the function of mitotic protein(s) of interest can be studied effectively, even without the use of live cell imaging.
Figure 2 shows that licensing of DNA replication origin is not perturbed when cells are depleted of Cdt1 during G2/M phase using our protocol, which was also observed in control cells. Cells depleted of Cdt1 during G2/M phase did not induce accumulation of phosphorylated &#611;H2AX, a marker for DNA damage, during the subsequent G2 phase; whereas siRNA transfection of asynchronous cultures to deplete Cdt1 induced accumulation of phospho-&#611;H2AX-positive foci, possibly due to DNA damage induced by improper DNA replication licensing.
Study of mitotic progression in cells after release from double thymidine block by live cell imaging 
After the nuclear envelope breakdown (NEB), normal cells (in the absence of perturbation by a functional mitotic protein) enter mitosis and undergo anaphase onset to exit from mitosis within about 30-60 min (Figure 3B). But RNAi-mediated knockdown of Hec1, a key kinetochore protein that is required for robust kMT attachment formation, is found to delay normal mitotic progression. In this scenario, most cells enter mitosis after the NEB, but do not undergo anaphase onset or exit from mitosis even after several hours of delay in mitosis (Figure 3C). Similarly, the effect of RNAi-mediated knockdown of any other protein that localizes to kinetochores or having a function during mitosis can thus be observed effectively by using live cell imaging after release from double thymidine blocks. In addition to studying the nature of mitotic progression, delay, and arrest, some of the other key mitotic phenomena that can be assayed using this method include chromosome alignment, mitotic spindle formation, and positioning the activation and silencing of the spindle assembly checkpoint.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56513/56513fig1.jpg" /> 
Figure 1: Analysis of mitotic progression and the stability of kinetochore microtubules in fixed cells after double thymidine synchronization. (A) A schematic representation of cell synchronization, fixation, and immunofluorescence. (B) Western blot for knockdown of Cdt1 after 9 h of release from 2nd thymidine block. (C) Immunofluorescence micrographs to show the mitotic cells fixed 9 and 10 h after release from double thymidine block and treatment with control (top) or Cdt1 siRNA (bottom). &#945;-tubulin staining is in red, DNA is in green as cells express GFP-Histone H2B. Scale bar = 10 &#181;m. Prometaphase and metaphase cells are indicated using yellow and white arrowheads respectively. (D) HeLa cells after treatment with control siRNA (top panel) and after Cdt1 depletion (bottom panel) for 9 h after release from double thymidine block followed by treatment with ice-cold Leibovitz&#39;s (L-15) medium and fixation. Cells were immune-stained for spindle MT (green) and a kinetochore marker, Zwint1 (red) and DAPI for DNA in black and white. Scale bar = 5 &#181;m. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56513/56513fig1large.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/56513/56513fig2.jpg" /> 
Figure 2: Cdt1-depletion during G2/M phase does not perturb replication licensing. Asynchronous HeLa cells treated with control luciferase (top panel) or cdt1 siRNAs (middle panel) or double thymidine synchronized HeLa cells treated with cdt1 siRNA during the 2nd thymidine wash-out followed by fixation at 10 h after release (bottom panel) were stained with DAPI (pseudo-colored green) and anti-phospho-&#611;H2AX antibody (red). Scale Bar = 10 &#181;m. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56513/56513fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56513/56513fig3.jpg" /> 
Figure 3: Analysis of mitotic progression by live cell imaging in synchronized cells. (A) A schematic representation of cell synchronization and live cell imaging. HeLa cells stably expressing mCherry-Histone H2B and GFP-&#945;-tubulin were released from double thymidine block for 8 h to let them proceed to G2/M phase from S phase arrest. Live imaging was carried out using a high resolution confocal microscope starting 8 h after releasing the cells from thymidine block and continuing for up to 16 h from that point. Shown here are the still images from the video captured every 10 min for control cells without perturbation of any mitotic protein (B) or for cells depleted of the Hec1 subunit of the Ndc80 complex (C). Scale Bars = 10 &#181;m. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56513/56513fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Combining Mitotic Cell Synchronization and High Resolution Confocal Microscopy to Study the Role of Multifunctional Cell Cycle Proteins During Mitosis at JoVE

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

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 ...

combining-mitotic-cell-synchronization-high-resolution-confocal

Universidad San Sebastian

Research

JoVE Journal

Biology

14.2K Views.  Northwestern University. 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. 

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

Studying Cell Cycle-regulated Gene Expression by Two Complementary Cell Synchronization Protocols

The gene expression program of the cell cycle represents a critical step for understanding cell cycle-dependent processes and their role in diseases such as cancer. Cell cycle-regulated gene expression analysis depends on cell synchronization into specific phases. Here we describe a method utilizing two complementary synchronization protocols that is commonly used for studying periodic variation of gene expression during the cell cycle. Both procedures are based on transiently blocking the cell cycle in one defined point. The synchronization protocol by hydroxyurea (HU) treatment leads to cellular arrest in late G1/early S phase, and release from HU-mediated arrest provides a cellular population uniformly progressing through S and G2/M. The synchronization protocol by thymidine and nocodazole (Thy-Noc) treatment blocks cells in early mitosis, and release from Thy-Noc mediated arrest provides a synchronized cellular population suitable for G1 phase and S phase-entry studies. Application of both procedures requires monitoring of the cell cycle distribution profiles, which is typically performed after propidium iodide (PI) staining of the cells and flow cytometry-mediated analysis of DNA content. We show that the combined use of two synchronization protocols is a robust approach to clearly determine the transcriptional profiles of genes that are differentially regulated in the cell cycle (i.e. E2F1 and E2F7), and consequently to have a better understanding of their role in cell cycle processes. Furthermore, we show that this approach is useful for the study of mechanisms underlying drug-based therapies (i.e. mitomycin C, an anticancer agent), because it allows to discriminate genes that are responsive to the genotoxic agent from those solely affected by cell cycle perturbations imposed by the agent.

We report two cell synchronization protocols that provide a context for studying events related to specific phases of the cell cycle. We show that this approach is useful for analyzing the regulation of specific genes in an unperturbed cell cycle or upon exposure to agents affecting the cell cycle.

The gene expression program of the cell cycle represents a critical step for understanding cell cycle-dependent processes and their role in diseases such ...

Genetics

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.

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 ...

Developmental Biology

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 ...

Mammalian Cell Division in 3D Matrices via Quantitative Confocal Reflection Microscopy

The study of how mammalian cell division is regulated in a 3D environment remains largely unexplored despite its physiological relevance and therapeutic significance. Possible reasons for the lack of exploration are the experimental limitations and technical challenges that render the study of cell division in 3D culture inefficient. Here, we describe an imaging-based method to efficiently study mammalian cell division and cell-matrix interactions in 3D collagen matrices. Cells labeled with fluorescent H2B are synchronized using the combination of thymidine blocking and nocodazole treatment, followed by a mechanical shake-off technique. Synchronized cells are then embedded into a 3D collagen matrix. Cell division is monitored using live-cell microscopy. The deformation of collagen fibers during and after cell division, which is an indicator of cell-matrix interaction, can be monitored and quantified using quantitative confocal reflection microscopy. The method provides an efficient and general approach to study mammalian cell division and cell-matrix interactions in a physiologically relevant 3D environment. This approach not only provides novel insights into the molecular basis of the development of normal tissue and diseases, but also allows for the design of novel diagnostic and therapeutic approaches.

This protocol efficiently studies mammalian cell division in 3D collagen matrices by integrating synchronization of cell division, monitoring of division events in 3D matrices using live-cell imaging technique, time-resolved confocal reflection microscopy and quantitative imaging analysis.

The study of how mammalian cell division is regulated in a 3D environment remains largely unexplored despite its physiological relevance and therapeutic ...

Bioengineering

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 ...

A Faster, High Resolution, mtPA-GFP-based Mitochondrial Fusion Assay Acquiring Kinetic Data of Multiple Cells in Parallel Using Confocal Microscopy

Mitochondrial fusion plays an essential role in mitochondrial calcium homeostasis, bioenergetics, autophagy and quality control. Fusion is quantified in living cells by photo-conversion of matrix targeted photoactivatable GFP (mtPAGFP) in a subset of mitochondria. The rate at which the photoconverted molecules equilibrate across the entire mitochondrial population is used as a measure of fusion activity. Thus far measurements were performed using a single cell time lapse approach, quantifying the equilibration in one cell over an hour. Here, we scale up and automate a previously published live cell method based on using mtPAGFP and a low concentration of TMRE (15 nm). This method involves photoactivating a small portion of the mitochondrial network, collecting highly resolved stacks of confocal sections every 15 min for 1 hour, and quantifying the change in signal intensity. Depending on several factors such as ease of finding PAGFP expressing cells, and the signal of the photoactivated regions, it is possible to collect around 10 cells within the 15 min intervals. This provides a significant improvement in the time efficiency of this assay while maintaining the highly resolved subcellular quantification as well as the kinetic parameters necessary to capture the detail of mitochondrial behavior in its native cytoarchitectural environment.
Mitochondrial dynamics play a role in many cellular processes including respiration, calcium regulation, and apoptosis1,2,3,13. The structure of the mitochondrial network affects the function of mitochondria, and the way they interact with the rest of the cell. Undergoing constant division and fusion, mitochondrial networks attain various shapes ranging from highly fused networks, to being more fragmented. Interestingly, Alzheimer's disease, Parkinson's disease, Charcot Marie Tooth 2A, and dominant optic atrophy have been correlated with altered mitochondrial morphology, namely fragmented networks4,10,13. Often times, upon fragmentation, mitochondria become depolarized, and upon accumulation this leads to impaired cell function18. Mitochondrial fission has been shown to signal a cell to progress toward apoptosis. It can also provide a mechanism by which to separate depolarized and inactive mitochondria to keep the bulk of the network robust14. Fusion of mitochondria, on the other hand, leads to sharing of matrix proteins, solutes, mtDNA and the electrochemical gradient, and also seems to prevent progression to apoptosis9. How fission and fusion of mitochondria affects cell homeostasis and ultimately the functioning of the organism needs further understanding, and therefore the continuous development and optimization of how to gather information on these phenomena is necessary.
Existing mitochondrial fusion assays have revealed various insights into mitochondrial physiology, each having its own advantages. The hybrid PEG fusion assay7, mixes two populations of differently labeled cells (mtRFP and mtYFP), and analyzes the amount of mixing and colocalization of fluorophores in fused, multinucleated, cells. Although this method has yielded valuable information, not all cell types can fuse, and the conditions under which fusion is stimulated involves the use of toxic drugs that likely affect the normal fusion process. More recently, a cell free technique has been devised, using isolated mitochondria to observe fusion events based on a luciferase assay1,5. Two human cell lines are targeted with either the amino or a carboxy terminal part of Renilla luciferase along with a leucine zipper to ensure dimerization upon mixing. Mitochondria are isolated from each cell line, and fused. The fusion reaction can occur without the cytosol under physiological conditions in the presence of energy, appropriate temperature and inner mitochondrial membrane potential. Interestingly, the cytosol was found to modulate the extent of fusion, demonstrating that cell signaling regulates the fusion process 4,5. This assay will be very useful for high throughput screening to identify components of the fusion machinery and also pharmacological compounds that may affect mitochondrial dynamics. However, more detailed whole cell mitochondrial assays will be needed to complement this in vitro assay to observe these events within a cellular environment.
A technique for monitoring whole-cell mitochondrial dynamics has been in use for some time and is based on a mitochondrially-targeted photoactivatable GFP (mtPAGFP)6,11. Upon expression of the mtPAGFP, a small portion of the mitochondrial network is photoactivated (10-20%), and the spread of the signal to the rest of the mitochondrial network is recorded every 15 minutes for 1 hour using time lapse confocal imaging. Each fusion event leads to a dilution of signal intensity, enabling quantification of the fusion rate. Although fusion and fission are continuously occurring in cells, this technique only monitors fusion as fission does not lead to a dilution of the PAGFP signal6. Co-labeling with low levels of TMRE (7-15 nM in INS1 cells) allows quantification of the membrane potential of mitochondria. When mitochondria are hyperpolarized they uptake more TMRE, and when they depolarize they lose the TMRE dye. Mitochondria that depolarize no longer have a sufficient membrane potential and tend not to fuse as efficiently if at all. Therefore, active fusing mitochondria can be tracked with these low levels of TMRE9,15. Accumulation of depolarized mitochondria that lack a TMRE signal may be a sign of phototoxicity or cell death. Higher concentrations of TMRE render mitochondria very sensitive to laser light, and therefore great care must be taken to avoid overlabeling with TMRE. If the effect of depolarization of mitochondria is the topic of interest, a technique using slightly higher levels of TMRE and more intense laser light can be used to depolarize mitochondria in a controlled fashion (Mitra and Lippincott-Schwartz, 2010). To ensure that toxicity due to TMRE is not an issue, we suggest exposing loaded cells (3-15 nM TMRE) to the imaging parameters that will be used in the assay (perhaps 7 stacks of 6 optical sections in a row), and assessing cell health after 2 hours. If the mitochondria appear too fragmented and cells are dying, other mitochondrial markers, such as dsRED or Mitotracker red could be used instead of TMRE.
The mtPAGFP method has revealed details about mitochondrial network behavior that could not be visualized using other methods. For example, we now know that mitochondrial fusion can be full or transient, where matrix content can mix without changing the overall network morphology. Additionally, we know that the probability of fusion is independent of contact duration and organelle dimension, is influenced by organelle motility, membrane potential and history of previous fusion activity8,15,16,17.
In this manuscript, we describe a methodology for scaling up the previously published protocol using mtPAGFP and 15nM TMRE8 in order to examine multiple cells at a time and improve the time efficiency of data collection without sacrificing the subcellular resolution. This has been made possible by the use of an automated microscope stage, and programmable image acquisition software. Zen software from Zeiss allows the user to mark and track several designated cells expressing mtPAGFP. Each of these cells can be photoactivated in a particular region of interest, and stacks of confocal slices can be monitored for mtPAGFP signal as well as TMRE at specified intervals. Other confocal systems could be used to perform this protocol provided there is an automated stage that is programmable, an incubator with CO2, and a means by which to photoactivate the PAGFP; either a multiphoton laser, or a 405 nm diode laser.

Mitochondrial fusion was measured by tracking the equilibration of photoconverted matrix-targeted GFP across the mitochondrial network over time. Thus far, only one cell could be subjected to an hour long kinetic analysis at a time. We present a method that simultaneously measures multiple cells, thereby speeding up the data collection process.

Mitochondrial fusion plays an essential role in mitochondrial calcium homeostasis, bioenergetics, autophagy and quality control. Fusion is quantified in ...

Studying Proteolysis of Cyclin B at the Single Cell Level in Whole Cell Populations

Equal distribution of chromosomes between the two daughter cells during cell division is a prerequisite for guaranteeing genetic stability 1. Inaccuracies during chromosome separation are a hallmark of malignancy and associated with progressive disease 2-4. The spindle assembly checkpoint (SAC) is a mitotic surveillance mechanism that holds back cells at metaphase until every single chromosome has established a stable bipolar attachment to the mitotic spindle1. The SAC exerts its function by interference with the activating APC/C subunit Cdc20 to block proteolysis of securin and cyclin B and thus chromosome separation and mitotic exit. Improper attachment of chromosomes prevents silencing of SAC signaling and causes continued inhibition of APC/CCdc20 until the problem is solved to avoid chromosome missegregation, aneuploidy and malignant growths1.
Most studies that addressed the influence of improper chromosomal attachment on APC/C-dependent proteolysis took advantage of spindle disruption using depolymerizing or microtubule-stabilizing drugs to interfere with chromosomal attachment to microtubules. Since interference with microtubule kinetics can affect the transport and localization of critical regulators, these procedures bear a risk of inducing artificial effects 5.
To study how the SAC interferes with APC/C-dependent proteolysis of cyclin B during mitosis in unperturbed cell populations, we established a histone H2-GFP-based system which allowed the simultaneous monitoring of metaphase alignment of mitotic chromosomes and proteolysis of cyclin B 6.
To depict proteolytic profiles, we generated a chimeric cyclin B reporter molecule with a C-terminal SNAP moiety 6 (Figure 1). In a self-labeling reaction, the SNAP-moiety is able to form covalent bonds with alkylguanine-carriers (SNAP substrate) 7,8 (Figure 1). SNAP substrate molecules are readily available and carry a broad spectrum of different fluorochromes. Chimeric cyclin B-SNAP molecules become labeled upon addition of the membrane-permeable SNAP substrate to the growth medium 7 (Figure 1). Following the labeling reaction, the cyclin B-SNAP fluorescence intensity drops in a pulse-chase reaction-like manner and fluorescence intensities reflect levels of cyclin B degradation 6 (Figure 1). Our system facilitates the monitoring of mitotic APC/C-dependent proteolysis in large numbers of cells (or several cell populations) in parallel. Thereby, the system may be a valuable tool to identify agents/small molecules that are able to interfere with proteolytic activity at the metaphase to anaphase transition. Moreover, as synthesis of cyclin B during mitosis has recently been suggested as an important mechanism in fostering a mitotic block in mice and humans by keeping cyclin B expression levels stable 9,10, this system enabled us to analyze cyclin B proteolysis as one element of a balanced equilibrium 6.

Metaphase to anaphase transition is triggered through anaphase-promoting complex (APC/C)-dependent ubiquitination and subsequent destruction of cyclin B. Here, we established a system which, following pulse-chase labeling, allows monitoring cyclin B proteolysis in entire cell populations and facilitates the detection of interference by the mitotic checkpoint.

Equal distribution of chromosomes between the two daughter cells during cell division is a prerequisite for guaranteeing genetic stability 1. Inaccuracies ...

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, ...

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

Summary

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Chapters in this video

Studying Cell Cycle-regulated Gene Expression by Two Complementary Cell Synchronization Protocols

Live Cell Imaging of Chromosome Segregation During Mitosis

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Time-lapse Imaging of Mitosis After siRNA Transfection

Mammalian Cell Division in 3D Matrices via Quantitative Confocal Reflection Microscopy

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

A Faster, High Resolution, mtPA-GFP-based Mitochondrial Fusion Assay Acquiring Kinetic Data of Multiple Cells in Parallel Using Confocal Microscopy

Studying Proteolysis of Cyclin B at the Single Cell Level in Whole Cell Populations

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