The authors would like to thank the efforts of Palak Sekhri, Hussam Alkhalaileh, Hsiaochi Chang, and Justin Lyeberger for their experimental support. This work was supported by the Pelotonia Fellowship Program. Any opinions, findings, and conclusions expressed in this material are those of the author(s) and do not necessarily reflect those of the Pelotonia Fellowship Program.&#34;

1. Preparation of IdU stocks
<ol>
	<li>Dissolve 5-iodo-2&#8217;-deoxyuridine (IdU) in DMSO to a concentration of 50 mM. Sterile filter, aliquot into 10-50 &#181;L tubes, and store at -80 &#176;C</li>
	<li>Remove IdU from the freezer, and thaw at room temperature. Dilute IdU in RPMI-1640 to make a working solution at a final concentration of 1 mM. Pipette up and down or vortex to mix.
	<ol>
		<li>Typically, dilute the concentrated IdU into the media in which the cells are being cultured (e.g. DMEM, IMDM, etc.) or have diluted it into PBS for addition directly to peripheral blood or bone marrow aspirate samples. This pre-dilution step facilitates mixing of the DMSO with the aqueous media of the cells of interest. 
		NOTE: The final concentration of IdU during incubation should be 10 &#181;M; a solution of 1 mM will can be added at a ratio of 10 &#181;L to every 1 mL of media.</li>
	</ol>
	</li>
</ol>
2. IdU incubation and sample preservation
<ol>
	<li>Maintain samples in a humidified 37 &#176;C incubator. Remove the sample from the incubator and move the sample into a biosafety hood.</li>
	<li>Add 10 &#181;L of 1 mM IdU to every 1 mL of sample.
	<ol>
		<li>For a 6-well plate, add 30 &#181;L of 1 mM IdU to the 3 mL of culture media in each well. IdU can also be used directly in bone marrow aspirates as well as murine studies4.</li>
	</ol>
	</li>
	<li>Place the sample back into the incubator at 37 &#176;C for 10-15 min. Maintain cells under the optimal growth conditions of interest during IdU exposure in order to get the most accurate measurement of S-phase.</li>
	<li>After the IdU incubation, remove the sample and transfer to a conical tube.</li>
	<li>Spin the sample at 400 x g for 10 min at room temperature.</li>
	<li>Aspirate the supernatant and resuspend in 200 &#181;L of PBS.
	<ol>
		<li>If needed, perform a live/dead stain (using cisplatin) at this step to mark dead cells before fixation and freezing. 
		NOTE: Rhodium live/dead staining does not perform well after methanol permeabilization, so it is not recommended for use in cell cycle analyses.</li>
	</ol>
	</li>
	<li>Add 18.75 &#181;L of 16% paraformaldehyde (PFA) to the PBS for a final concentration of 1.5% PFA. Incubate at room temperature for 10 min. Spin the sample down at 400 x g for 10 min at room temperature.</li>
	<li>Aspirate the PBS/PFA solution and re-suspend the sample in 500 &#181;L of Cell Staining Media (CSM; 1x PBS with 0.5% BSA and 0.02% sodium azide) + 10% DMSO prior to freezing.
	<ol>
		<li>If using commercial Proteomic Stabilizer, add 280 &#181;L of Proteomic Stabilizer to sample re-suspended in 200 &#181;L of PBS (1:1.4). Incubate samples at room temperature for 10 min and then place directly into the -80 &#176;C. 
		NOTE: IdU has been shown to incorporate effectively within 10-15 min incubation at 37 &#176;C. IdU incubations longer than 10-15 min will progressively reduce resolution of the S and G2-phase populations, as IdU-labeled cells leave S phase and progress to G2 or M phase. We have also observed that long-term incubation with IdU can cause cell death and cell cycle artifacts. The subsequent cell processing and antibody staining following IdU incorporation is sufficient to wash away residual IdU that was not incorporated into S-phase cells. We have not observed significant Iodine background when using the in vitro protocol described here; however, we have very rarely observed iodine contamination in clinical samples. This may occur from medical procedures, such as iodine contrast in a CT scan, or from iodine-containing pharmaceuticals. Should large amounts of IdU background be observed, the sample should not be run to avoid damage to the mass cytometer&#8217;s detector.</li>
	</ol>
	</li>
</ol>
3. Staining samples for mass cytometry
<ol>
	<li>Remove the samples from the -80 &#176;C and allow to thaw before surface staining.
	<ol>
		<li>If using the SmartTube method of fixation, thaw the samples at 0-4 &#176;C to avoid additional fixation as the samples warm up.</li>
	</ol>
	</li>
	<li>After the samples are thawed, transfer approximately 1-2 million cells into a 5 mL FACS tube.</li>
	<li>Centrifuge the FACS tube at 600 x g for 5 min, and fill the FACS tube with cell staining media (CSM) to wash the cells. Repeat one additional time.
	<ol>
		<li>If cells are known to stick together, add 400 U/mL of heparin to CSM washes in order to prevent cell to cell contact but this is not strictly necessary.</li>
	</ol>
	</li>
	<li>Incubate the cells with FC-blocking agent, 5 &#181;L of the agent per 100 &#181;L of cells, for 10 min at room temperature.</li>
	<li>Prepare a mixture of antibodies that will stain the surface, or extracellular portion, of the cells. The total staining mixture will amount to 100 &#181;L per 1-2 million cells in each test. The staining mixture will be balanced accordingly with CSM and heparin in the cocktail and FACS tube. 
	NOTE: Addition of CSM at this step will also reduce nonspecific staining artifacts8. This staining mixture is entirely dependent on targets of interest and surface phenotype (e.g., a study involving T-cells will use a surface mixture of CD45, CD3, CD4, CD8, etc.). A detailed protocol of sample processing and staining can be found in Behbehani and McCarthy et al.9,10.</li>
	<li>Add the surface staining mixture to the cells and incubate at room temperature with continuous shaking for 30-60 min.</li>
	<li>After staining, fill the FACS tube with CSM, and spin down at 600 x g for 5 min.</li>
	<li>Wash two more times with CSM, spinning the sample at 600 x g for 5 min and aspirating each CSM wash.</li>
	<li>Fix the extracellular antibodies by adding 1 mL of PBS with 10% CSM and 1.5% PFA.</li>
	<li>Fill the FACS tube containing the PBS/CSM/PFA mixture with CSM. Spin down at 600 x g for 5 min and aspirate the supernatant.</li>
	<li>Add methanol at -20 &#176;C.</li>
	<li>Vortex the sample for 1-2 min to achieve a single cell suspension and verify that all cell clumps have been re-suspended.</li>
	<li>While the sample is vortexing slowly, rapidly add 1 mL of ice-cold methanol using a 1,000 &#181;L pipette with a filter tip.</li>
	<li>Hold the FACs tube up to the light and make sure there are no visible clumps; cloudiness is to be expected. Any clumps will render the sample unusable for subsequent MCM analysis.</li>
	<li>Store the sample at -20 &#176;C for 10-20 min.</li>
	<li>Prepare the intracellular staining mixture during this time. The intracellular staining mixture will be dependent on targets of interest. For cell cycle analysis, include CyclinB1, pRb, Ki67, and pHH3 in this staining mixture, but other intracellular markers can be added as needed.</li>
	<li>After 10-20 min at -20 &#176;C, remove the sample, add 1.5 mL of PBS and fill the remainder with CSM.</li>
	<li>Centrifuge the sample 600 x g for 5 min, and aspirate the supernatant.</li>
	<li>Wash two more times with CSM, spinning the sample down at 600 x g for 5 min and aspirating the supernatant each time.</li>
	<li>After the last CSM wash, centrifuge the sample and leave a residual volume of approximately 50 &#181;L.</li>
	<li>Add the prepared antibody mixture (typically add 50 &#181;L of antibody staining cocktail to achieve a final staining volume of 100 &#181;L) to the sample and incubate on a shaking platform for 30 to 60 min at room temperature.</li>
	<li>After staining add CSM and centrifuge at 600 x g for 5 min.</li>
	<li>Aspirate the CSM, wash again with CSM, spinning at 600 x g for 5 min, aspirating the CSM, then add PBS.</li>
	<li>After the completion of intracellular staining, place cells into an intercalator solution that fixes the antibodies to the cells and stains the DNA of each cells to enable identification. The intercalator solution contains nonisotopically pure iridium intercalator (pentamethylcyclopentadienyl-Ir(III)-dipyridophenazine) added from the manufacturer&#8217;s stock solution at a concentration of 500 &#181;M. Dilute the Iridium stock 1:4000 in a solution of PBS and 1.5% PFA. Add the iridium intercalator solution at 100-200 &#181;L per million cells in order to stain evenly and prevent overstaining. 
	NOTE: The iridium in this intercalator solution is intended to identify cells for singlet gating, it should not be used for live/dead stains. If live/dead stains are desired they need to be performed before fixation as noted above and in McCarthy et al.9.</li>
	<li>Store the samples in intercalator solution in a 4 &#176;C refrigerator for up to two weeks before sample acquisition on the CyTOF.</li>
</ol>
4. Mass cytometer operation
NOTE: Mass cytometry operation can be machine specific. It is always advisable to check the CyTOF user&#8217;s manual before operation. Additionally, there are currently two JoVE articles dealing with machine start up and maintenance9,11.
<ol>
	<li>Check the nebulizer for any clogs, cracks, and other irregularities before operating the mass cytometer.</li>
	<li>Connect the nebulizer to the cytometer and begin the warmup procedure. Do not start the mass cytometer without the nebulizer in place.</li>
	<li>Run water through the sample lines once the mass cytometer has finished warming up. The spray chamber needs to reach approximately 200 &#176;C before performing tuning or sample analysis.</li>
	<li>Run water for 5-10 min. After 5-10 min, load the tuning solution and select the tuning manager. The tuning solution is a solution containing fixed concentrations of metals and used to optimize the mass cytometer before sample acquisition</li>
	<li>In the tuning manager, select Preview once the tuning solution has reached a steady state hit record to begin the automated tuning process.</li>
	<li>Once tuning is finished load the sampler with water and allow water to run through the sample lines during sample processing. Detailed protocol for daily cytometer operation and tuning can be found at Leipold11.</li>
	<li>Occasionally, the automated tuning will not tune to optimal machine performance. Repeat the tuning procedure to correct this.</li>
	<li>Wash the sample with CSM once and with pure deionized water twice before sample acquisition. Washing with water is important to remove residual salt from the PBS/CSM.</li>
	<li>Check the sensitivity and sample flow using manufacturer supplied equilibration beads, polystyrene beads loaded with known metal concentrations.</li>
	<li>Change the acquisition mode from tuning to event capture mode. Set the time limit to stop acquisition at 120 s. Wait 45 s before selecting Record.</li>
	<li>The mass cytometer will stop sample acquisition automatically after 120 seconds. Use the rain plot viewer to check Eu151 and Eu153 intensity.</li>
	<li>Dilute equilibration beads in pure deionized water at a 1:20 ratio.</li>
	<li>Before sample acquisition check the experiment manager. Use the experiment manager to assign names to channels and to add channels to be recorded.
	<ol>
		<li>Make sure the 127-I channel is added if using IdU.</li>
		<li>Note that it is essential to set the mass cytometer to measure the needed parameters (e.g., IdU) prior to sample acquisition. If channels are not selected in advance, data will not be collected from any unselected channel and cannot be recovered.</li>
	</ol>
	</li>
	<li>Dilute the cells to a concentration of approximately 1-2 x 106/mL using the 1:20 pure deionized water and equilibration bead mixture. Pass the cells through the filter topped FACS tube in order to remove any residual clumps.</li>
	<li>Load the sample and change the acquisition time.</li>
	<li>Press Preview and wait for event count per second to stabilize.
	<ol>
		<li>Do not run events in excess of 400 events per second, this will lead to significant amounts of doublets and debris. We typically collect at least 20,000 to 50,000 cell events, but the optimal number will depend on the experimental design. Staining of up to 2 million cells will typically yield 300,000 to 400,000 cell events. Note that not all events will be cells (there will be debris and bead events included in the event count).</li>
	</ol>
	</li>
	<li>Once sample acquisition is done load a washing solution, start sample induction and run for 5-10 minutes. After 5-10 minutes stop sample induction and run water for 10-20 minutes. Wash solution is a weak solution of hydrofluoric acid designed to strip residual metal from the sample lines.</li>
	<li>Shut down the mass cytometer and remove the nebulizer. The nebulizer will be hot, take care during handling.</li>
</ol>
5. Data analysis
<ol>
	<li>In order to remove beads and also to correct for signal drift during sample acquisition, normalize the FCS files using Fluidigm software or the application developed by Finck12.</li>
	<li>Upload the FCS to Cytobank or other flow cytometry analysis software. FCS files can be used in any compatible software, for the purposes of this protocol all gating and further analysis has been done in Cytobank13.</li>
	<li>Before cell cycle gates can be drawn, exclude any doublets or cell debris from downstream analysis, this can be done by using the biaxial plot of Event Length vs 191-Ir (Figure 1a). Cells will form a distinct, bright population Irhigh that can be used to exclude doublets and debris. This is the singlet gate. This gating method typically removes about 50-60% of doublet cell events, so additional strategies may be required to remove remaining doublet cell events.
	<ol>
		<li>Change the event length scale (minimum and maximum) to make the cells appear more prominent to aid in singlet gating.</li>
		<li>Further remove doublets and debris by using Gaussian parameters, Residual and Offset. A higher Residual with lower Offset is also debris and doublets, and gating around this population can further remove doublets and debris (Figure 1b,c).</li>
	</ol>
	</li>
	<li>S-phase gating &#8211; S-phase is the easiest gate to draw but also the most important. Draw this gate using a biaxial plot of IdU vs pRb, Ki67, or cyclinB1. S-phase IdU+ cells will form a distinct population when looking at these biaxial plots (Figure 2b).</li>
	<li>G0/G1-phase, G2/M-phase gating &#8211; Establish the G0/G1 and G2/M phase gates on the IdU vs CyclinB1 plot and the use of IdU incorporation is crucial to establish the boundary between the G0/G1 and G2/M phase gates. G0/G1-phase will be CyclinB1low/IdU- and G2/M-phase will be CyclinB1high/IdU-. Good CyclinB1 staining will show a natural population between the G0-G1 and G2-M populations; however, this will vary across sample and cell types under experimental conditions. In experimental conditions where the cell cycle distribution may be affected and there is less separation between the CyclinB1 G0/G1-phase and G2/M-phase utilizing the S-phase will allow consistent gating for particular cell types in each specific experiment. This method is detailed below.
	<ol>
		<li>Plot only the S-phase cells on the CyclinB1 vs IdU to help establish the separation between G0/G1-phase and G2/M-phase. Draw a gate on the CyclinB1high population and adjust until approximately the top 5% of the S-phase population is inside the gate (Figure 2c). This establishes the breakpoint between the G0/G1 and G2/M phase gates (Figure 2e,f). The active population will be changed to the population of interest and the portion residing inside the previous gate will be the G2/M-phase population while the remainder will be the G0/G1-phase population.</li>
	</ol>
	</li>
	<li>G0-phase gating &#8211; Establish the G0-phase on the pRb vs IdU plot. The G0-phase will be represented by a pRblow/IdU- population. The active cycling population will have high expression of pRb and IdU incorporation, the G0-phase gate can be drawn on this boundary as it typically expresses at two distinct populations (Figure 2g,i).
	<ol>
		<li>Define the G0-phase by making the S-phase population drawn previously (Figure 2b) the active population and drawing a gate incorporating the top 90-100% of the pRbhigh population. This is the pRb+ cycling population (Figure 2i), the pRblow population outside this gate is the G0 population (Figure 2j).</li>
		<li>If pRb is unavailable or not able to be recorded, use Ki67 vs IdU to establish the G0-phase population. Drawing a gate representing the majority of the S-phase population and using that gate as the boundary for the Ki67 vs IdU the remainder of the population will be the G0-phase (Figure 3a).</li>
	</ol>
	</li>
	<li>M-phase gating &#8211; Establish the M-phase in the IdU vs pH3 biaxial plot. The M-phase represents a very small fraction of cells and is gated on the pH3high population (Figure 2d).</li>
	<li>IdU incorporation failed or was not possible &#8211; If IdU is unavailable, define cell cycling and not cycling fractions using Ki67 and pRb. Ki67 and pRb, in normal conditions, form two distinct populations a Ki67high/pRbhigh and a Ki67low/pRblow. The double positive population represents the active cycling population, correlating to G1-phase, S-phase, G2-phase, and M-phase. The double low population represents the not cycling population, correlating to the G0-phase (Figure 3b). 
	NOTE: It is not possible to delineate each individual phase using the Ki67 vs pRb but experimental effects on the relative cycling/not cycling populations can be determined.</li>
	<li>Cell cycle analysis &#8211; Once the gates have been established, export the numerical values from the gates for further analysis. The percentages in each cycle can be achieved by subtracting the single populations from the combined populations. The G0-phase, drawn on the pRblowIdUlow, gate percentage can be subtracted from the G0/G1-phase, drawn on CyclinB1lowIdUneg, to find the G1-phase percentage. Similarly the G2-phase percentage is derived from the subtraction of the M-phase gate from the G2/M-phase gate. This will generate numerical values for each individual cell cycle phase; G0, G1, S, G2, and M. The numerical values generated for each individual cell cycle phase can be used for further analysis such as graphing and statistical analysis.</li>
</ol>

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	<li>Ornatsky, O., Baranov, V. I., Bandura, D. R., Tanner, S. D., Dick, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiple+cellular+antigen+detection+by+ICP-MS.">Multiple cellular antigen detection by ICP-MS.</a> Journal of Immunological Methods. 308 (1-2), 68-76 (2006).</li><li>Ornatsky, O. I., 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=Study+of+cell+antigens+and+intracellular+DNA+by+identification+of+element-containing+labels+and+metallointercalators+using+inductively+coupled+plasma+mass+spectrometry.">Study of cell antigens and intracellular DNA by identification of element-containing labels and metallointercalators using inductively coupled plasma mass spectrometry.</a> Analytical Chemistry. 80 (7), 2539-2547 (2008).</li><li>Behbehani, G. K., Bendall, S. C., Clutter, M. 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Dr. Behbehani receives travel support from Fluidigm. Fluidigm has also purchased reagents and materials for lab use.

The examples presented here demonstrate how to use an MCM platform to analyze cell cycle distribution. It has also been demonstrated that cell cycle analysis is sensitive to experimental conditions such as time and temperature, which is an important consideration researchers must take when considering MCM for their cell cycle analysis14. Samples left in storage for a short period of time, no longer than an hour, will have IdU incorporation comparable to their normal state. Samples in a closed syst...

Mass cytometry enables detection of approximately 40 parameters by taking advantage of the high resolution and quantitative nature of mass spectroscopy. Metal-labeled antibodies are used instead of fluorophore conjugated antibodies that allow for a higher number of channels and produce minimal spillover1,2. MCM has advantages and disadvantages in regard to cell cycle analysis in comparison to flow cytometry. One major advantage of MCM is that the large number of parameters enables the simultaneous measurement of cell cycle state across a large number of immunophenotypically distinct T-cell types in highly heterogeneous samples. MCM has been successfully used to measure the cell cycle state during normal hematopoiesis in human bone marrow3 and transgenic murine models of telomerase deficiency4. Analysis of cell cycle state in acute myeloid leukemia (AML) showed that cell cycle correlated to known responses to clinical therapies, providing an in vivo insight into functional characteristics that can inform therapy selections5. A second advantage of mass cytometric cell cycle analysis is the ability to measure a large number of other functional markers that may be correlated with cell cycle state. Recent work has been able to correlate protein and RNA synthesis with cell cycle state through the use of IdU and metal tagged antibodies to BRU and rRNA6. This kind of highly parametric analysis measuring cell cycle state across numerous populations in a continuum of differentiation would be nearly impossible with current flow cytometry technology. The major disadvantage of MCM is the lack of comparable DNA or RNA stains as those used in fluorescent flow cytometry (e.g., DAPI, Hoechst, Pyronin Y, etc.). Fluorescent dyes can give relatively precise measurements of DNA and RNA content, but this precision is only possible due to the changes in the fluorescent properties of these dyes that occur upon intercalation between nucleotide bases. MCM analysis is thus unable to measure DNA or RNA content with similar precision. Instead, mass cytometric cell cycle analysis relies on measurements of proteins related to cell cycle state such as cyclin B1, phosphorylated retinoblastoma protein (pRb), and phosphorylated Histone H3 (pHH3) combined with direct measurement of the iodine atom from IdU incorporation into S-phase cells. These two measurement approaches yield highly similar results during normal cellular proliferation, but can potentially be discordant when cell cycle progression is disrupted.
Measurement of the number of cells in each cell cycle phase is important in understanding normal cell cycle development as well as cell cycle disruption, which is commonly observed in cancers and immunological diseases. MCM provides reliable measurement of extracellular and intracellular factors using metal-tagged antibodies; however, measurement of the S-phase was limited as the iridium-based DNA intercalator was unable to differentiate between 2N and 4N DNA. In order to define cell cycle phases, Behbehani developed a method that utilizes IdU with a mass of 127, which falls within the range of the mass cytometer and allows direct measurement of cells in S-phase3. This direct measurement circumvents the need for secondary antibodies or use of DNA denaturing agents such as acid or DNase. In conjunction with intracellular cycling markers, it allows high resolution of cell cycle distribution in experimental models.
This protocol adapts cell cycle measurements from common flow cytometry protocols for MCM. Our methods provide a convenient and simple way to include cell cycle parameters. IdU incorporation of in vitro samples requires only 10 to 15 minutes of incubation at 37 &#176;C, which is shorter than most BrdU staining protocols that recommend incubation times of several hours3,7. IdU and BrdU incorporated samples can be fixed using a proteomic stabilizer and then stored for some time in a -80 &#176;C freezer. This allows large numbers of IdU stained samples to be archived for batch analysis without reduction in sample quality.

<table><tbody><tr><td>Bovine Serum Albumin (BSA)</td><td>Sigma</td><td>A3059</td><td>Component of CSM</td></tr><tr><td>Centrifuge</td><td>Thermo Scientific</td><td>75-217-420</td><td>Sample centrifugation</td></tr><tr><td>Cleaved-PARP (D214)</td><td>BD Biosciences</td><td>F21-852</td><td>Identification of apoptotic cells</td></tr><tr><td>Cyclin B1</td><td>BD Biosciences</td><td>GNS-1</td><td>G2 Resolution</td></tr><tr><td>Dimethylsulfoxide (DMSO)</td><td>Sigma</td><td>D2650</td><td>Cryopreservative</td></tr><tr><td>EQ Four Element Calibration Beads</td><td>Fluidigm</td><td>201078</td><td>Internal metal standard for CyTOF performance</td></tr><tr><td>FACS Tube w/ mesh strainer</td><td>Corning</td><td>08-771-23</td><td>Cell strainer to remove clumps/debris before CyTOF run</td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>VWR</td><td>97068-085</td><td>Cell culture growth supplement</td></tr><tr><td>Helios</td><td>Fluidigm</td><td></td><td>CyTOF System/Platform</td></tr><tr><td>Heparin</td><td>Sigma</td><td>H3393</td><td>Staining additive to prevent non-specific staining</td></tr><tr><td>IdU (5-Iodo-2&amp;prime;-deoxyuridine)</td><td>Sigma</td><td>I7125</td><td>Incorporates in S-phase</td></tr><tr><td>Ki-67</td><td>eBiosciences</td><td>SolA15</td><td>Confirmation of G0/G1</td></tr><tr><td>MaxPar Multi Label Kit</td><td>Fluidigm</td><td>201300</td><td>Metal labeling kit, attaches metals to antibodies</td></tr><tr><td>Microplate Shaker</td><td>Thermo Scientific</td><td>88880023</td><td>Mixing samples during staining</td></tr><tr><td>Paraformaldehyde (PFA)</td><td>Electron Microscopy Services</td><td>15710</td><td>Fixative</td></tr><tr><td>pentamethylcyclopentadienyl-Ir(III)-dipyridophenazine</td><td>Fluidigm</td><td>201192</td><td>Cell identification during CyTOF acquisition</td></tr><tr><td>p-H2AX (S139)</td><td>Millipore</td><td>JBW301</td><td>Detection of DNA damage</td></tr><tr><td>p-HH3 (S28)</td><td>Biolegend</td><td>HTA28</td><td>M-phase Resolution</td></tr><tr><td>Phosphate Buffered Saline (PBS)</td><td>Gibco</td><td>14190-144</td><td>Wash solution for cell culture and component of fixative solution</td></tr><tr><td>p-Rb (S807/811)</td><td>BD Biosciences</td><td>J112906</td><td>G0/G1 Resolution</td></tr><tr><td>Proteomic Stabilizer</td><td>SmartTube Inc</td><td>PROT1</td><td>Sample fixative</td></tr><tr><td>RPMI 1640</td><td>Gibco</td><td>21870-076</td><td>Cell culture growth medium</td></tr><tr><td>Sodium Azide</td><td>Acros Organics</td><td>AC447810250</td><td>Component of CSM/Antibody buffer, biocide</td></tr></tbody></table>

use of the pyrimidine analog, 5-iodo-2&prime;-deoxyuridine (idu) with cell cycle markers to establish cell cycle phases in a mass cytometry platform

The regulation of cell cycle phase is an important aspect of cellular proliferation and homeostasis. Disruption of the regulatory mechanisms governing the cell cycle is a feature of a number of diseases, including cancer. Study of the cell cycle necessitates the ability to define the number of cells in each portion of cell cycle progression as well as to clearly delineate between each cell cycle phase. The advent of mass cytometry (MCM) provides tremendous potential for high throughput single cell analysis through direct measurements of elemental isotopes, and the development of a method to measure the cell cycle state by MCM further extends the utility of MCM. Here we describe a method that directly measures 5-iodo-2&#8242;-deoxyuridine (IdU), similar to 5-bromo-2&#180;-deoxyuridine (BrdU), in an MCM system. Use of this IdU-based MCM provides several advantages. First, IdU is rapidly incorporated into DNA during its synthesis, allowing reliable measurement of cells in the S-phase with incubations as short as 10-15 minutes. Second, IdU is measured without the need for secondary antibodies or the need for DNA degradation. Third, IdU staining can be easily combined with measurement of cyclin B1, phosphorylated retinoblastoma protein (pRb), and phosphorylated histone H3 (pHH3), which collectively provides clear delineation of the five cell cycle phases. Combination of these cell cycle markers with the high number of parameters possible with MCM allow combination with numerous other metrics.

Utilizing HL-60 cells and a human bone marrow aspirate it is possible to show how experimental conditions can affect cell cycle distribution and analysis. First, the gating strategy must be established to demonstrate how the cell cycle phases are derived. In Figure 1 we show the establishment of the singlet gate, which is important in separating cellular debris and doublets, establishing a single cell population. For cell lines the singlet gate is all that is needed to move onto cell cycle a...

Watch this Scientific Journal Video about Use of the Pyrimidine Analog, 5-Iodo-2â€²-Deoxyuridine IdU with Cell Cycle Markers to Establish Cell Cycle Phases in a Mass Cytometry Platform at JoVE.com

Use of the Pyrimidine Analog, 5-Iodo-2&#8242;-Deoxyuridine IdU with Cell Cycle Markers to Establish Cell Cycle Phases in a Mass Cytometry Platform

This protocol adapts cell cycle measurements for use in a mass cytometry platform. With the multi-parameter capabilities of mass cytometry, direct measurement of iodine incorporation allows identification of cells in S-phase while intracellular cycling markers enable characterization of each cell cycle state in a range of experimental conditions.

Use of the Pyrimidine Analog, 5-Iodo-2&prime;-Deoxyuridine (IdU) with Cell Cycle Markers to Establish Cell Cycle Phases in a Mass Cytometry Platform

The regulation of cell cycle phase is an important aspect of cellular proliferation and homeostasis. Disruption of the regulatory mechanisms governing the ...

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Genetics

2.5K Views.  The Ohio State University. This protocol adapts cell cycle measurements for use in a mass cytometry platform. With the multi-parameter capabilities of mass cytometry, direct measurement of iodine incorporation allows identification of cells in S-phase while intracellular cycling markers enable characterization of each cell cycle state in a range of experimental conditions.

Video: Use of the Pyrimidine Analog, 5-Iodo-2′-Deoxyuridine IdU with Cell Cycle Markers to Establish Cell Cycle Phases in a Mass Cytometry Platform

Isolation and Staining of Mouse Skin Keratinocytes for Cell Cycle Specific Analysis of Cellular Protein Expression by Mass Cytometry

The goal of this protocol is to detect and quantify protein expression changes in a cell cycle-dependent manner using single cells isolated from the epidermis of mouse skin. There are seven important steps: separation of the epidermis from the dermis, digestion of the epidermis, staining of the epidermal cell populations with cisplatin, sample barcoding, staining with metal tagged antibodies for cell cycle markers and proteins of interest, detection of metal-tagged antibodies by mass cytometry, and the analysis of expression in the various cell cycle phases. The advantage of this approach over histological methods is the potential to assay the expression pattern of &#62;40 different markers in a single cell at different phases of the cell cycle. This approach also allows for the multivariate correlation analysis of protein expression that is more quantifiable than histological/imaging methods. The disadvantage of this protocol is that a suspension of single cells is needed, which results in the loss of location information provided by the staining of tissue sections. This approach may also require the inclusion of additional markers to identify different cell types in crude cell suspensions. The application of this protocol is evident in the analysis of hyperplastic skin disease models. Moreover, this protocol can be adapted for the analysis of specific sub-type of cells (e.g., stem cells) by the addition of lineage-specific antibodies. This protocol can also be adapted for the analysis of skin cells in other experimental species.

This protocol describes how to isolate skin keratinocytes from mouse models, to stain with metal-tagged antibodies, and to analyze stained cells by mass cytometry in order to profile the expression pattern of proteins of interest in the different cell cycle phases.

The goal of this protocol is to detect and quantify protein expression changes in a cell cycle-dependent manner using single cells isolated from the ...

Developmental Biology

Immunofluorescent Detection of Two Thymidine Analogues (CldU and IdU) in Primary Tissue

Accurate measurement of cell division is a fundamental challenge in experimental biology that becomes increasingly complex when slowly dividing cells are analyzed. Established methods to detect cell division include direct visualization by continuous microscopy in cell culture, dilution of vital dyes such as carboxy&#64258;uorescein di-aetate succinimidyl ester (CFSE), immuno-detection of mitogenic antigens such as ki67 or PCNA, and thymidine analogues. Thymidine analogues can be detected by a variety of methods including radio-detection for tritiated thymidine, immuno-detection for bromo-deoxyuridine (BrdU), chloro-deoxyuridine (CldU) and iodo-deoxyuridine (IdU), and chemical detection for ethinyl-deoxyuridine (EdU). We have derived a strategy to detect sequential incorporation of different thymidine analogues (CldU and IdU) into tissues of adult mice. Our method allows investigators to accurately quantify two successive rounds of cell division. By optimizing immunostaining protocols our approach can detect very low dose thymidine analogues administered via the drinking water, safe to administer to mice for prolonged periods of time. Consequently, our technique can be used to detect cell turnover in very long-lived tissues. Optimal immunofluoresent staining results can be achieved in multiple tissue types, including pancreas, skin, gut, liver, adrenal, testis, ovary, thyroid, lymph node, and brain. We have also applied this technique to identify oncogenic transformation within tissues. We have further applied this technique to determine if transit-amplifying cells contribute to growth or renewal of tissues. In this sense, sequential administration of thymidine analogues represents a novel approach for studying the origins and survival of cells involved in tissue homeostasis.

Immunofluorescent Detection of Two Thymidine Analogues CldU and IdU in Primary Tissue

We have derived a strategy to detect sequential incorporation of thymidine analogues (CldU and IdU) into tissues of adult mice to quantify two successive rounds of cell division. This strategy is useful to detect cell turnover of long-lived tissues, oncogenic transformation, or transit-amplifying cells.

Accurate measurement of cell division is a fundamental challenge in experimental biology that becomes increasingly complex when slowly dividing cells are ...

Biology

Analysis of Cell Cycle Position in Mammalian Cells

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of cell proliferation underlies the pathology of diseases like cancer. As such there is great need to be able to investigate cell proliferation and quantitate the proportion of cells in each phase of the cell cycle. It is also of vital importance to indistinguishably identify cells that are replicating their DNA within a larger population. Since a cell&prime;s decision to proliferate is made in the G1 phase immediately before initiating DNA synthesis and progressing through the rest of the cell cycle, detection of DNA synthesis at this stage allows for an unambiguous determination of the status of growth regulation in cell culture experiments.
DNA content in cells can be readily quantitated by flow cytometry of cells stained with propidium iodide, a fluorescent DNA intercalating dye. Similarly, active DNA synthesis can be quantitated by culturing cells in the presence of radioactive thymidine, harvesting the cells, and measuring the incorporation of radioactivity into an acid insoluble fraction. We have considerable expertise with cell cycle analysis and recommend a different approach. We Investigate cell proliferation using bromodeoxyuridine/fluorodeoxyuridine (abbreviated simply as BrdU) staining that detects the incorporation of these thymine analogs into recently synthesized DNA. Labeling and staining cells with BrdU, combined with total DNA staining by propidium iodide and analysis by flow cytometry1 offers the most accurate measure of cells in the various stages of the cell cycle. It is our preferred method because it combines the detection of active DNA synthesis, through antibody based staining of BrdU, with total DNA content from propidium iodide. This allows for the clear separation of cells in G1 from early S phase, or late S phase from G2/M. Furthermore, this approach can be utilized to investigate the effects of many different cell stimuli and pharmacologic agents on the regulation of progression through these different cell cycle phases.
In this report we describe methods for labeling and staining cultured cells, as well as their analysis by flow cytometry. We also include experimental examples of how this method can be used to measure the effects of growth inhibiting signals from cytokines such as TGF-&beta;1, and proliferative inhibitors such as the cyclin dependent kinase inhibitor, p27KIP1. We also include an alternate protocol that allows for the analysis of cell cycle position in a sub-population of cells within a larger culture5. In this case, we demonstrate how to detect a cell cycle arrest in cells transfected with the retinoblastoma gene even when greatly outnumbered by untransfected cells in the same culture. These examples illustrate the many ways that DNA staining and flow cytometry can be utilized and adapted to investigate fundamental questions of mammalian cell cycle control.

Determining the cell cycle position of a population of cells, or understanding how signals affect proliferation, can be readily measured by flow cytometry using this protocol. We report a simple experimental approach to staining cells and quantifying their position in the cell cycle.

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of ...

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating cells. Cell cycle division is an integral part of growth and reproduction and deregulation of key cell cycle components have been implicated in the precipitating events of carcinogenesis 1,2. Molecular agents in anti-cancer therapies frequently target biological pathways responsible for the regulation and coordination of cell cycle division 3. Although cell cycle kinetics tend to vary according to cell type, the distribution of cells amongst the four stages of the cell cycle is rather consistent within a particular cell line due to the consistent pattern of mitogen and growth factor expression. Genotoxic events and other cellular stressors can result in a temporary block of cell cycle progression, resulting in arrest or a temporary pause in a particular cell cycle phase to allow for instigation of the appropriate response mechanism. 
The ability to experimentally observe the behavior of a cell population with reference to their cell cycle progression stage is an important advance in cell biology. Common procedures such as mitotic shake off, differential centrifugation or flow cytometry-based sorting are used to isolate cells at specific stages of the cell cycle 4-6. These fractionated, cell cycle phase-enriched populations are then subjected to experimental treatments. Yield, purity and viability of the separated fractions can often be compromised using these physical separation methods. As well, the time lapse between separation of the cell populations and the start of experimental treatment, whereby the fractionated cells can progress from the selected cell cycle stage, can pose significant challenges in the successful implementation and interpretation of these experiments.
Other approaches to study cell cycle stages include the use of chemicals to synchronize cells. Treatment of cells with chemical inhibitors of key metabolic processes for each cell cycle stage are useful in blocking the progression of the cell cycle to the next stage. For example, the ribonucleotide reductase inhibitor hydroxyurea halts cells at the G1/S juncture by limiting the supply of deoxynucleotides, the building blocks of DNA. Other notable chemicals include treatment with aphidicolin, a polymerase alpha inhibitor for G1 arrest, treatment with colchicine and nocodazole, both of which interfere with mitotic spindle formation to halt cells in M phase and finally, treatment with the DNA chain terminator 5-fluorodeoxyridine to initiate S phase arrest 7-9. Treatment with these chemicals is an effective means of synchronizing an entire population of cells at a particular phase. With removal of the chemical, cells rejoin the cell cycle in unison. Treatment of the test agent following release from the cell cycle blocking chemical ensures that the drug response elicited is from a uniform, cell cycle stage-specific population. However, since many of the chemical synchronizers are known genotoxic compounds, teasing apart the participation of various response pathways (to the synchronizers vs. the test agents) is challenging. 
Here we describe a metabolic labeling method for following a subpopulation of actively cycling cells through their progression from the DNA replication phase, through to the division and separation of their daughter cells. Coupled with flow cytometry quantification, this protocol enables for measurement of kinetic progression of the cell cycle in the absence of either mechanically- or chemically- induced cellular stresses commonly associated with other cell cycle synchronization methodologies 10. In the following sections we will discuss the methodology, as well as some of its applications in biomedical research.

Tracking subtle changes in the progression and kinetics of cell cycle stages can be accomplished by use of a combination of metabolic labeling of nucleic acids with BrdU and total genomic DNA staining via Propidium Iodide. This method avoids the need of chemical synchronization of cycling cells, thereby preventing the introduction of non-specific DNA damage, which in turn affects cell cycle progression.

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating ...

Temporal Tracking of Cell Cycle Progression Using Flow Cytometry without the Need for Synchronization

This protocol describes a method to permit the tracking of cells through the cell cycle without requiring the cells to be synchronized. Achieving cell synchronization can be difficult for many cell systems. Standard practice is to block cell cycle progression at a specific stage and then release the accumulated cells producing a wave of cells progressing through the cycle in unison. However, some cell types find this block toxic resulting in abnormal cell cycling, or even mass death. Bromodeoxyuridine (BrdU) uptake can be used to track the cell cycle stage of individual cells. Cells incorporate this synthetic thymidine analog, while synthesizing new DNA during S phase. By providing BrdU for a brief period it is possible to mark a pool of cells that were in S phase while the BrdU was present. These cells can then be tracked through the remainder of the cell cycle and into the next round of replication, permitting the duration of the cell cycle phases to be determined without the need to induce a potentially toxic cell cycle block. It is also possible to determine and correlate the expression of both internal and external proteins during subsequent stages of the cell cycle. These can be used to further refine the assignment of cell cycle stage or assess effects on other cellular functions such as checkpoint activation or cell death.

This protocol describes the use of bromodeoxyuridine (BrdU) uptake to permit the temporal tracking of cells that were in S phase at a specific point in time. Addition of DNA dyes and antibody labeling facilitates detailed analysis of the fate of the S phase cells at later times.

This protocol describes a method to permit the tracking of cells through the cell cycle without requiring the cells to be synchronized. Achieving cell ...

Experimental Approaches to Study Mitochondrial Localization and Function of a Nuclear Cell Cycle Kinase, Cdk1

Although mitochondria possess their own transcriptional machinery, merely 1% of mitochondrial proteins are synthesized inside the organelle. The nuclear-encoded proteins are transported into mitochondria guided by their mitochondria targeting sequences (MTS); however, a majority of mitochondrial localized proteins lack an identifiable MTS. Nevertheless, the fact that MTS can instruct proteins to go into the mitochondria provides a valuable tool for studying mitochondrial functions of normally nuclear and/or cytoplasmic proteins. We have recently identified the cell cycle kinase CyclinB1/Cdk1 complex in the mitochondria. To specifically study the mitochondrial functions of this complex, mitochondrial overexpression and knock-down of this complex without interfering with its nuclear or cytoplasmic functions were essential. By tagging CyclinB1/Cdk1 with MTS, we were able to achieve mitochondrial overexpression of this complex to study its mitochondrial targets as well as functions. Via tagging dominant-negative Cdk1 with MTS, inhibition of Cdk1 activity was accomplished particularly in the mitochondria. Potential mitochondrial targets of CyclinB1/Cdk1 complex were identified using a gel-based proteomics approach. Unlike traditional 2D gel analysis, we employed 2-dimensional difference gel electrophoresis (2D-DIGE) technology followed by phosphoprotein staining to fluorescently label differentially phosphorylated proteins in mitochondrial Cdk1 expressing cells. Identification of phosphoprotein spots that were altered in wild type versus dominant negative Cdk1 bearing mitochondria revealed the identity of mitochondrial targets of Cdk1. Finally, to determine the effect of CyclinB1/Cdk1 mitochondrial localization in cell cycle progression, a cell proliferation assay using a synthetic thymidine analogue EdU (5-ethynyl-2&#8242;-deoxyuridine) was used to monitor the cells as they go through the cell cycle and replicate their DNA. Altogether, we demonstrated a variety of approaches available to study mitochondrial localization and activity of a cell cycle kinase. These are advanced, yet easy to follow methods that will be beneficial to many cell biology researchers.

Here, we outline how to study mitochondrial localization of a (cell cycle) kinase, and how to determine its sub-mitochondrial location as well as potential mitochondrial substrates/targets. Forced expression of proteins into the mitochondria provides a useful tool for studying the functional consequences of mitochondrial localization of a protein of interest.

Although mitochondria possess their own transcriptional machinery, merely 1% of mitochondrial proteins are synthesized inside the organelle. The ...

Assessment of Global DNA Double-Strand End Resection using BrdU-DNA Labeling coupled with Cell Cycle Discrimination Imaging

The study of the DNA damage response (DDR) is a complex and essential field, which has only become more important due to the use of DDR-targeting drugs for cancer treatment. These targets are poly(ADP-ribose) polymerases (PARPs), which initiate various forms of DNA repair. Inhibiting these enzymes using PARP inhibitors (PARPi) achieves synthetic lethality by conferring a therapeutic vulnerability in homologous recombination (HR)-deficient cells due to mutations in breast cancer type 1 (BRCA1), BRCA2, or partner and localizer of BRCA2 (PALB2). 
Cells treated with PARPi accumulate DNA double-strand breaks (DSBs). These breaks are processed by the DNA end resection machinery, leading to the formation of single-stranded (ss) DNA and subsequent DNA repair. In a BRCA1-deficient context, reinvigorating DNA resection through mutations in DNA resection inhibitors, such as 53BP1 and DYNLL1, causes PARPi resistance. Therefore, being able to monitor DNA resection in cellulo is critical for a clearer understanding of the DNA repair pathways and the development of new strategies to overcome PARPi resistance. Immunofluorescence (IF)-based techniques allow for monitoring of global DNA resection after DNA damage. This strategy requires long-pulse genomic DNA labeling with 5-bromo-2&#8242;-deoxyuridine (BrdU). Following DNA damage and DNA end resection, the resulting single-stranded DNA is specifically detected by an anti-BrdU antibody under native conditions. Moreover, DNA resection can also be studied using cell cycle markers to differentiate between various phases of the cell cycle. Cells in the S/G2 phase allow the study of end resection within HR, whereas G1 cells can be used to study non-homologous end joining (NHEJ). A detailed protocol for this IF method coupled to cell cycle discrimination is described in this paper.

In the present protocol, we demonstrate how to visualize DNA double-strand end resection during S/G2 phase of the cell cycle using an immunofluorescence-based method.

The study of the DNA damage response (DDR) is a complex and essential field, which has only become more important due to the use of DDR-targeting drugs ...

5-Ethynyl-2'-Deoxyuridine/Phospho-Histone H3 Dual-Labeling Protocol for Cell Cycle Progression Analysis in Drosophila Neural Stem Cells

In vivo cell cycle progression analysis is routinely performed in studies on genes regulating mitosis and DNA replication. 5-Ethynyl-2'-deoxyuridine (EdU) has been utilized to investigate replicative/S-phase progression, whereas antibodies against phospho-histone H3 have been utilized to mark mitotic nuclei and cells. A combination of both labels would enable the classification of G0/G1 (Gap phase), S (replicative), and M (mitotic) phases and serve as an important tool to evaluate the effects of mitotic gene knockdowns or null mutants on cell cycle progression. However, the reagents used to mark EdU-labelled cells are incompatible with several secondary antibody-fluorescent tags. This complicates immunostaining, where primary and tagged secondary antibodies are used to mark pH3-positive mitotic cells. This paper describes a step-by-step protocol for the dual-labeling of EdU and pH3 in Drosophila larval neural stem cells, a system utilized extensively to study mitotic factors. Additionally, a protocol is provided for image analysis and quantification to allocate labeled cells in 3 distinct categories, G0/G1, S, S&gt;G2/M (progression from S to G2/M), and M phases.

5-Ethynyl-2'-Deoxyuridine/Phospho-Histone H3 Dual-Labeling Protocol for Cell Cycle Progression Analysis in Drosophila Neural Stem Cells

Cell cycle analysis with 5-ethynyl-2'-deoxyuridine (EdU) and phospho-histone H3 (pH3) labeling is a multi-step procedure that may require extensive optimization. Here, we present a detailed protocol that describes all steps for this procedure including image analysis and quantification to distinguish cells in different cell cycle phases.

In vivo cell cycle progression analysis is routinely performed in studies on genes regulating mitosis and DNA replication. 5-Ethynyl-2'-deoxyuridine (EdU) ...

Determination of S-Phase Duration Using 5-Ethynyl-2'-deoxyuridine Incorporation in Saccharomyces cerevisiae

Eukaryotic DNA replication is a highly regulated process that ensures that the genetic blueprint of a cell is correctly duplicated prior to chromosome segregation. As DNA synthesis defects underlie chromosome rearrangements, monitoring DNA replication has become essential to understand the basis of genome instability. Saccharomyces cerevisiae is a classical model to study cell cycle regulation, but key DNA replication parameters, such as the fraction of cells in the S phase or the S-phase duration, are still difficult to determine. This protocol uses short and non-toxic pulses of 5-ethynyl-2'-deoxyuridine (EdU), a thymidine analog, in engineered TK-hENT1 yeast cells, followed by its detection by Click reaction to allow the visualization and quantification of DNA replication with high spatial and temporal resolution at both the single-cell and population levels by microscopy and flow cytometry. This method may identify previously overlooked defects in the S phase and cell cycle progression of yeast mutants, thereby allowing the characterization of new players essential for ensuring genome stability.

Determination of S-Phase Duration Using 5-Ethynyl-2'-deoxyuridine Incorporation in Saccharomyces cerevisiae

We describe two complementary protocols to accurately determine S-phase duration in S. cerevisiae using EdU, a thymidine analog, which is incorporated in vivo and detected using Click chemistry by microscopy and flow cytometry. It allows for the easy characterization of the duration of DNA replication and overlooked replication defects in mutants.

Eukaryotic DNA replication is a highly regulated process that ensures that the genetic blueprint of a cell is correctly duplicated prior to chromosome ...

Cell Cycle Analysis

Cell cycle refers to the set of events through which a cell grows, replicates its genome, and ultimately divides into two daughter cells through the process of mitosis. Because the amount of DNA in a cell shows characteristic changes throughout the cycle, techniques known as cell cycle analysis can be used to separate a population of cells according to the different phases of cell cycle they are in, based on their varying DNA content.This video will cover the principles behind cell cycle analysis via DNA-staining. We will review a generalized protocol for performing this staining using bromodeoxyuridine (BrdU, a thymidine analog that is incorporated into newly synthesized DNA strands) and propidium iodide (PI, a DNA dye that stains all DNA), followed by analysis of the stained cells with flow cytometry. During flow cytometry, a single cell suspension of fluorescently labeled cells is passed through an instrument with a laser beam and the fluorescence of each cell is read. We will then discuss how to interpret data from flow cytometric scatter plots, and finally, look at a few applications of this technique.

Cell Cycle Analysis: Principle, BrdU and Flow Cytometry

Cell cycle refers to the set of events through which a cell grows, replicates its genome, and ultimately divides into two daughter cells through the ...

Use of the Pyrimidine Analog, 5-Iodo-2′-Deoxyuridine (IdU) with Cell Cycle Markers to Establish Cell Cycle Phases in a Mass Cytometry Platform

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Chapters in this video

Isolation and Staining of Mouse Skin Keratinocytes for Cell Cycle Specific Analysis of Cellular Protein Expression by Mass Cytometry

Immunofluorescent Detection of Two Thymidine Analogues (CldU and IdU) in Primary Tissue

Analysis of Cell Cycle Position in Mammalian Cells

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

Temporal Tracking of Cell Cycle Progression Using Flow Cytometry without the Need for Synchronization

Experimental Approaches to Study Mitochondrial Localization and Function of a Nuclear Cell Cycle Kinase, Cdk1

Assessment of Global DNA Double-Strand End Resection using BrdU-DNA Labeling coupled with Cell Cycle Discrimination Imaging

5-Ethynyl-2'-Deoxyuridine/Phospho-Histone H3 Dual-Labeling Protocol for Cell Cycle Progression Analysis in Drosophila Neural Stem Cells

Determination of S-Phase Duration Using 5-Ethynyl-2'-deoxyuridine Incorporation in Saccharomyces cerevisiae

Cell Cycle Analysis