The work was funded by the Leukemia and Lymphoma Society of the USA (6105-08), a Cancer Council NSW grant (13-02), an NHMRC Senior Research Fellowship (LJB) (1042305) and project grant (1041614).

The protocol described here uses the acute lymphoblastic leukemia cell line NALM6 but can be applied to other cell types.
1. Solutions and Reagents
<ol>
	<li>Complete RPMI
	<ol>
		<li>Add 56 ml fetal calf serum (FCS) and 5.5 ml of 200 mM L-glutamine to a 500 ml bottle of RPMI-1640 medium.</li>
	</ol>
	</li>
	<li>BrdU Stock Solution
	<ol>
		<li>Prepare 32.5 mM BrdU (10 mg/ml) in Dulbecco&#39;s Phosphate Buffered Saline (DPBS).</li>
	</ol>
	</li>
	<li>BrdU Complete RPMI
	<ol>
		<li>Add 6.2 &#181;l of BrdU stock solution to 10 ml of Complete RPMI.</li>
	</ol>
	</li>
	<li>DNase Solution
	<ol>
		<li>Prepare 1 mg DNase/ml in DPBS.</li>
	</ol>
	</li>
	<li>Staining Buffer
	<ol>
		<li>Prepare 3% heat-inactivated FCS and 0.09% sodium azide in DPBS.</li>
	</ol>
	</li>
	<li>Refer to Materials List for definitions of Fixation Buffer, Permeabilization Buffer and Wash Buffer.</li>
</ol>
2. Cells
Note: Cells were not cultured for greater than 6 months. This method is directly adaptable to any non-adherent cell line with adjustments to cell density and culture media. Use cells that are growing exponentially at the initiation of the experiment.
<ol>
	<li>Maintain NALM6 cells in T-75 culture flasks in Complete RPMI. Perform all steps under sterile conditions using a Class II Biosafety Cabinet.
	<ol>
		<li>Maintain NALM6 cells between 1-2 x 106 cells per ml by splitting the culture thrice weekly.</li>
		<li>Incubate at 37 &#176;C in 5% CO2 in air.</li>
	</ol>
	</li>
</ol>
3. Pulse Labeling of Cells with BrdU
CAUTION: Handle BrdU with care as it is a potential mutagen and teratogen.
<ol>
	<li>Centrifuge cells at 150 x g for 5 min. Note: Transferring cells into fresh media improves the reproducibility of the results.</li>
	<li>Perform a cell count and resuspend cells in Complete RPMI at 2 x 106 cells/ml.</li>
	<li>Dilute cells 1 in 2 with BrdU Complete RPMI producing a final cell concentration of 1 x 106 cells/ml.</li>
	<li>Incubate at 37 &#176;C with 5% CO2 for 45 min, then dilute cells 1 in 10 with complete RPMI. Centrifuge cells at 150 x g for 5 min and carefully discard all of the supernatant.</li>
	<li>Resuspend cells in a small volume (~100 &#181;l) of complete RPMI, perform a cell count and adjust to 1 x 106 cells/ml.</li>
	<li>Pipette 1 ml of cells into the wells of a 48 well plate. Pipette 1 ml of DPBS into any unoccupied wells to obtain more reproducible results.</li>
	<li>Incubate at 37 &#176;C in 5% CO2 in air for desired timepoints, here 1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, and 24 hr. Note: The length of time will depend on what the experimental design aims to measure.</li>
	<li>Transfer all the cells into FACS tubes using a pipette. Rinse the well sequentially with 1 ml volumes of PBS to a final total volume of 5 ml.</li>
	<li>Centrifuge at 150 x g for 5 min and carefully remove all the supernatant. Cells are ready for staining, perform this (section 4) immediately.</li>
</ol>
4. Cell Staining
Note: If surface staining of cells is required perform it prior to fixation, ensuring that the cells are kept at 4 &#176;C throughout.
<ol>
	<li>Resuspend cells in 100 &#181;l of staining buffer (for optional surface staining, add the recommended volume of antibody to surface antigens and incubate for 30 min at 4 &#176;C).</li>
	<li>Add 1 ml of staining buffer, centrifuge for 5 min at 150 x g and discard the supernatant. 
	Note: Specific antibody, concentration, incubation time etc. will vary depending on specific experimental goals.</li>
	<li>Fixation and Permeabilization
	<ol>
		<li>Resuspend cells in 100 &#181;l of fixation buffer and incubate for 15 min at room temperature.</li>
		<li>Add 1 ml of wash buffer, centrifuge for 5 min at 150 x g and discard the supernatant.</li>
		<li>Resuspend cells in 100 &#181;l of permeabilization buffer and incubate the cells for 10 min on ice.</li>
		<li>Add 1 ml of wash buffer, centrifuge for 5 min at 150 x g, and discard the supernatant.</li>
		<li>Resuspend cells in 100 &#181;l of fixation buffer per tube and incubate for 5 min at room temperature.</li>
		<li>Add 1 ml of wash buffer, centrifuge for 5 min at 150 x g, and discard the supernatant. 
		Note: The protocol can be paused here if required. The fixed cells are stable for several days at 4 &#176;C if resuspended in staining buffer. Remove the staining buffer following centrifugation before proceeding.</li>
	</ol>
	</li>
	<li>DNase Treatment
	<ol>
		<li>Resuspend cells in 100 &#181;l of DNase solution (30 &#181;g of DNase/106 cells) and incubate cells for 1 hr at 37 &#176;C.</li>
		<li>Add 1 ml of wash buffer, centrifuge at 150 x g for 5 min and discard supernatant.</li>
	</ol>
	</li>
	<li>Antibody Staining 
	Note: Staining for intracellular markers other than BrdU can be performed simultaneously with the BrdU staining.
	<ol>
		<li>IMPORTANT: Prepare compensation controls consisting of unstained cells and cells labeled with each single fluorochrome. Ideally, use the same antibodies for compensation controls as those used in the experimental tubes. However, if this is not feasible, substitute antibodies to highly expressed antigens conjugated to the same fluorochrome.</li>
		<li>Resuspend the cells in 50 &#181;l of wash buffer and add 1 &#181;l/106 cells of BrdU antibody. Note: Directly conjugated antibodies to other specific intracellular antigens can also be added. &#160; 
		NOTE: Antibodies to histone H3 phosphorylated on Ser10 can be used to discriminate between cells in G2 and M, histone H3 is phosphorylated on Ser10 during mitosis.10 Antibodies to cdc2 phosphorylated on Tyr15 can be used to detect cells that have committed to mitosis.11</li>
		<li>Incubate the cells for 20 min at room temperature.</li>
		<li>Add 1 ml of wash buffer, centrifuge cells at 150 x g for 5 min and discard supernatant.</li>
	</ol>
	</li>
</ol>
<ol start="6">
	<li>Stain DNA for Cell Cycle Analysis
	<ol>
		<li>Loosen pellet and add 20 &#181;l of the 7-AAD solution (0.25 &#181;g). Note: It is critical to use a constant amount of 7-AAD/cell.</li>
		<li>Resuspend the cells in 1 ml of Staining buffer.</li>
	</ol>
	</li>
</ol>
5. Collection of Flow Cytometry Data
Note: The machine required will depend on the number and nature of the fluorochromes used.
<ol>
	<li>Collect the following parameters: FSC-A, SSC-A, FSC-H (FSC-W can be used instead of FSC-H) and 7-AAD fluorescence on a linear scale. Collect the APC channel on a log scale. Collect any additional channels required for the assessment of surface or internal labels using a log scale.</li>
	<li>Perform compensation of overlapping signals in emission spectra observed between different fluorochromes before analyzing the samples. Note: Most flow cytometers will perform this automatically.</li>
	<li>Collect at least 10,000 events for each sample.</li>
</ol>
6. Analysis of Flow Cytometry Data
Note: FlowJo was used in this study for flow cytometry data analysis but other software packages can also be used. The gating strategy is illustrated in Figure 1.
<ol>
	<li>Identify the viable cell population using FSC-A and SSC-A parameters.</li>
	<li>Within this population exclude doublets and aggregates using FSC-A and FSC-H (FSC-W can also be used here).</li>
	<li>Within this population set a dot plot using 7-AAD on the x-axis and BrdU-APC on the y-axis.</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/52840/52840fig1highres.jpg" width="700" /> 
Figure 1: Gating Strategy. Left panel: ungated cells are shown on a FCS-A vs. SSC-A dot plot. The viable cell population is identified by the gate shown. Center panel: cells gated from the left panel are shown on a FSC-A vs. FSC-H dot plot (FSC-W can be used instead of height). Doublets and aggregates are identified, and excluded by the gate shown. Right panel: cells gated from the doublet exclusion date in the center panel are shown on a 7-AAD vs. APC-A dot plot. The BrdU antibody is labeled with APC permitting the identification of cells that have incorporated BrdU during the pulse labeling. 7-AAD provides information on DNA content. The upper gate defines cells positive for BrdU and therefore in S phase at the time of the BrdU pulse, the lower left gate, cells in G0/1 and the lower right gate those in G2/M. <a href="https://www.jove.com/files/ftp_upload/52840/52840fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="4">
	<li>Cell Cycle Analysis
	<ol>
		<li>Open the first data file and gate on the cells in the doublet exclusion gate.</li>
		<li>Analyze this population for cell cycle distribution (located under platforms in FloJo software) and use the Dean-Jett-Fox model.</li>
		<li>Obtain the positions of the G0/1 and G2/M peaks using create gates.</li>
		<li>Gate on the BrdU positive cells and subject this population to same cell cycle analysis.</li>
		<li>Provide the positions for the G0/1 and G2/M peaks by applying the same gates from create gates and setting constraints (using the created gates) for the positions of the G0/1 and G2/M peaks. This is illustrated in the first 2 panels of Figure 2. 
		Note: Other software may also be used to analyze the data and the instructions would vary accordingly.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/52840/52840fig2highres.jpg" width="700" /> 
Figure 2: Cell Cycle Progression. The first panel (All Cells) is gated on the cell population defined by the doublet exclusion gate. This population was displayed in a histogram with 7-AAD on the X-axis. The peak of the G0/1 peak is indicated by the arrow below the axis. In subsequent panels BrdU positive cells have been gated on as shown in Figure 1. The value for the G0/1 position obtained when gating of the doublet exclusion gate is applied to the BrdU positive gated cells within FlowJo cell cycle software. Each subsequent panel was gated on the BrdU positive population as shown in Figure 1 and the position of the G0/1 peak based on the value obtained when analyzing the whole population as shown in the first two panels. Using the BrdU negative fraction to identify the location of the G0/1 population for the BrdU positive cells in the same sample controls for any slight differences in the intensity of the DNA stain between samples. The number shown on each panel represents the time since the BrdU pulse ended. The calculated cell cycle phases are shown in shaded green. <a href="https://www.jove.com/files/ftp_upload/52840/52840fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors have nothing to disclose.

The ability to analyze the cell cycle is important for the understanding of cancer biology and the mechanism of action of both drugs and genes that influence cell proliferation and growth. While there are a multitude of assays that reportedly measure cell proliferation, the majority only provide a measure that indicates the number cells present. These include assays that measure cell number by direct visualization and counting, metabolic activity or ATP concentration. The main advantage of many of these methods is that t...

The assessment of cell cycle features and changes that occur in cells during cell cycle progression is fundamental to understanding many aspects of biology, particularly cancer biology. Many agents in development for the treatment of malignancies have profound effects on cell cycle progression or induce cell death via cell cycle dependent-mechanisms. In order to study cell cycle dynamics or cells in a particular phase of the cell cycle, it is usual to synchronize cells. However synchronization methods can have detrimental effects on the cells being studied, potentially confounding the results obtained.1 Recently the use of fluorescently tagged proteins that are only present at particular phases of the cells cycle have permitted analysis of cell cycle progression in single cells over time2, however the cells to be studied need to be genetically manipulated to express these tagged proteins, limiting their use to systems where this can be readily achieved.
The cell cycle consists of two active phases: the synthesis (S) phase, where DNA is replicated and mitosis (M) where cell division takes place. These phases are separated by three gap phases, G0, G1 and G2. G0 or quiescence, is a resting phase where the cell has left the cycle, G1 is where the cells increase in size prior to DNA replication and G2 where cell growth continues between completion of DNA replication but before cell division. The progression through the cell cycle is controlled by a number of checkpoints. The G1 checkpoint is activated when environmental conditions are not supportive of DNA synthesis and prevents entry into S phase. The intra-S phase checkpoint or delay can be triggered by DNA damage that may result in stalled replication forks. During G2 the fidelity of the replicated DNA is confirmed and if damage is detected then the G2 checkpoint is activated permitting DNA repair prior to cell division. A final checkpoint during mitosis ensures that chromatids have been correctly aligned at the mitotic plate so that cell division can be successfully completed.3 Activation of these checkpoints is commonly used to synchronize cell populations. Cell cycle checkpoints can be activated by a number of factors but in cancer biology the most common is detection of DNA damage. The DNA damage response is initiated by the PI3-kinase-like kinases ataxia telangiectasia and Rad3 related (ATR) and ataxia telangiectasia mutated (ATM) that activate the downstream effector kinases Chk1 and Chk2, respectively.3 A range of events activates Chk1 including stalled replication forks, DNA crosslinks, and ultraviolet radiation damage while Chk2 is primarily activated by double-strand breaks.
The usual method for studying the effect of altered conditions on the length of the cell cycle is to synchronize the cells in a particular phase of the cell cycle.1 This can be achieved via several methods. Cells can be physically separated based on size, density, side scatter (granularity), and cell surface expression markers. More practically, cells may be synchronized by chemical means. Several agents such as thymidine, hydroxyurea and cytosine arabinoside can be used to inhibit DNA synthesis in the S phase of cell cycle resulting in an accumulation of cells in S phase which continue cycling after the agents are removed. Cells treated with nocodazole, which prevents the formation of the mitotic spindle, arrest with a G2- or M-phase DNA content. Elimination of serum from the culture medium results in the accumulation of cells at G0 phase. The re-addition of the nutrients within the culture serum re-starts the normal cycling of the cells. However, all of these synchronization methods interfere with normal cycling and growth of cells and can result in significant cell death.
Synchronization of acute lymphoblastic leukemia cells is particularly challenging and these cells are not amenable to genetic manipulation. The method described here permits the assessment of cell cycle dynamics and the study of cells in particular phases of the cell cycle without traditional synchronization or genetic modification. This method may also be useful for other cell types where genetic modification and traditional synchronization procedures are not readily achieved. The method is based on the long established use of bromodeoxyuridine (BrdU) incorporation, which has very little impact on the short-term growth and proliferation of cells.4 Established BrdU protocols take advantage of the incorporation of BrdU into newly synthesized DNA during S phase. This permanently marks cells as having been in S phase during BrdU exposure. This population can be identified at later time points by staining for BrdU incorporation and thereby act as a synchronized population that can be followed and assessed over time permitting the study of drug effects on cell cycle transit. BrdU needs to be exposed prior to antibody staining, usually achieved following DNase or acid treatment.6,7 Using flow cytometry to detect incorporated BrdU enables the inclusion of additional markers. The most important is the use of dyes to measure DNA content, enabling the assessment of cell cycle phase distribution of the cells that were in S phase at the start of the study.8 Furthermore additional surface or intracellular antigens can also be studied.9 These may relate to cell cycle events such as Ki67 or to seemingly unrelated cell features such as apoptosis markers like cleaved caspase-3. The potential applications are limited by the imagination of the investigator.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>APC BrdU Flow Kit</td>
			<td>BD Biosciences</td>
			<td>552598</td>
			<td>Contains BrdU antibody, 7-AAD and BD Cytofix/Cytoperm 
			Buffer (referred to as Fixation Buffer)</td>
		</tr>
		<tr>
			<td>BD Cytoperm Permeabilization Buffer Plus</td>
			<td>BD Biosciences</td>
			<td>561651</td>
			<td>Referred to as Permeabilization buffer</td>
		</tr>
		<tr>
			<td>BD Perm/Wash Buffer</td>
			<td>BD Biosciences</td>
			<td>554723</td>
			<td>Referred to as Wash buffer</td>
		</tr>
		<tr>
			<td>DNase</td>
			<td>Sigma</td>
			<td>D-4513</td>
			<td></td>
		</tr>
		<tr>
			<td>BD Falcon 12 x 75-mm FACS tubes</td>
			<td>BD Biosciences</td>
			<td>352008</td>
			<td></td>
		</tr>
		<tr>
			<td>BD Pharmingen Stain Buffer</td>
			<td>BD Biosciences</td>
			<td>554656</td>
			<td></td>
		</tr>
		<tr>
			<td>BD LSR FORTESSA flow cytometer</td>
			<td>BD Biosciences</td>
			<td>FORTESSA</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipetman</td>
			<td>Gilson</td>
			<td>P2, P20, P100, P1000</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI 1640 w/o L-Gln 500 ml</td>
			<td>Lonza</td>
			<td>12-167F</td>
			<td></td>
		</tr>
		<tr>
			<td>DPBS</td>
			<td>Lonza</td>
			<td>17-512F</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>FisherBiotec</td>
			<td>FBS-7100113</td>
			<td></td>
		</tr>
		<tr>
			<td>L-Glutamine</td>
			<td>Sigma</td>
			<td>G7513-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>5-Bromo-2&#8242;-deoxyuridine</td>
			<td>Sigma</td>
			<td>B5002-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>Falcon TC 150cm2 vented Flasks</td>
			<td>BD Biosciences</td>
			<td>355001</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipettes 25mL</td>
			<td>Greiner</td>
			<td>760180</td>
			<td></td>
		</tr>
		<tr>
			<td>Aersol Pipettes 200&#181;L</td>
			<td>Interpath</td>
			<td>24700</td>
			<td></td>
		</tr>
		<tr>
			<td>Aersol Pipettes 1mL</td>
			<td>Interpath</td>
			<td>24800</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td>Spintron</td>
			<td>GT-175R</td>
			<td></td>
		</tr>
		<tr>
			<td>CO2 incubator</td>
			<td>Binder</td>
			<td>C 150</td>
			<td></td>
		</tr>
		<tr>
			<td>AF488 anti-Histone H3 Phospho (Ser10) Antibody</td>
			<td>Cell Signalling</td>
			<td>9708S</td>
			<td></td>
		</tr>
		<tr>
			<td>Phospho-Chk2 (Thr68) (C13C1) Rabbit mAb</td>
			<td>Cell Signalling</td>
			<td>2197S</td>
			<td></td>
		</tr>
		<tr>
			<td>Phospho-Chk1 (Ser345) (133D3) Rabbit mAb</td>
			<td>Cell Signalling</td>
			<td><a name="RANGE!C23">2348S</a></td>
			<td></td>
		</tr>
		<tr>
			<td>NALM6&#160;</td>
			<td>DSMZ</td>
			<td>ACC-128</td>
			<td></td>
		</tr>
	</tbody>
</table>

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 methodology can be used to obtain a range of information. A few applications are outlined here.
Assessment of the duration of the cell cycle
To determine the time required for cells to transit through the cell cycle, cells are harvested at various time points following the BrdU pulse. The intervals between assessments can be adapted to the particular cells being analyzed. Hematopoietic cell lines were assessed every hour over a 24 hr period in the absence of an...

Watch this Scientific Journal Video about Temporal Tracking of Cell Cycle Progression Using Flow Cytometry without the Need for Synchronization at JoVE.com

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

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 ...

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19.0K Views.  Westmead Millennium Institute for Medical Research and University of Sydney. 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.

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

Flow Cytometry Purification of Mouse Meiotic Cells

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the unique differentiation programs that are manifest during meiosis. Two principal methods have been developed to purify, to varying degrees, various meiotic fractions from both adult and immature animals: elutriation or Staput (sedimentation) using BSA and/or percoll gradients. Both of these methods rely on cell size and density to separate meiotic cells1-5. Overall, except for few cell populations6, these protocols fail to yield sufficient purity of the numerous meiotic cell populations that are necessary for detailed molecular analyses. Moreover, with such methods usually one type of meiotic cells can be purified at a given time, which adds an extra level of complexity regarding the reproducibility and homogeneity when comparing meiotic cell samples.
Here, we describe a refined method that allows one to easily visualize, identify, and purify meiotic cells, from germ cells to round spermatids, using FACS combined with Hoechst 33342 staining7,8. This method provides an overall snapshot of the entire meiotic process and allows one to highly purify viable cells from most stage of meiosis. These purified cells can then be analyzed in detail for molecular changes that accompany progression through meiosis, for example changes in gene expression9,10and the dynamics of nucleosome occupancy at hotspots of meiotic recombination11.

An efficient method to obtain highly purified viable meiotic fractions from mouse testis is described, which combines a refined cell dissociation protocol with fluorescent activated cell sorting (FACS). This method takes advantage of differences in the DNA content and nuclear density of discrete meiotic fractions.

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the ...

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 ...

Optimized Staining and Proliferation Modeling Methods for Cell Division Monitoring using Cell Tracking Dyes

Fluorescent cell tracking dyes, in combination with flow and image cytometry, are powerful tools with which to study the interactions and fates of different cell types in vitro and in vivo.1-5 Although there are literally thousands of publications using such dyes, some of the most commonly encountered cell tracking applications include monitoring of:
<ol>
	<li>stem and progenitor cell quiescence, proliferation and/or differentiation6-8</li>
	<li>antigen-driven membrane transfer9 and/or precursor cell proliferation3,4,10-18 and</li>
	<li>immune regulatory and effector cell function1,18-21.</li>
</ol>
Commercially available cell tracking dyes vary widely in their chemistries and fluorescence properties but the great majority fall into one of two classes based on their mechanism of cell labeling. "Membrane dyes", typified by PKH26, are highly lipophilic dyes that partition stably but non-covalently into cell membranes1,2,11. "Protein dyes", typified by CFSE, are amino-reactive dyes that form stable covalent bonds with cell proteins4,16,18. Each class has its own advantages and limitations. The key to their successful use, particularly in multicolor studies where multiple dyes are used to track different cell types, is therefore to understand the critical issues enabling optimal use of each class2-4,16,18,24.
The protocols included here highlight three common causes of poor or variable results when using cell-tracking dyes. These are:
<ol>
	<li>Failure to achieve bright, uniform, reproducible labeling. This is a necessary starting point for any cell tracking study but requires attention to different variables when using membrane dyes than when using protein dyes or equilibrium binding reagents such as antibodies.</li>
	<li>Suboptimal fluorochrome combinations and/or failure to include critical compensation controls. Tracking dye fluorescence is typically 102 - 103 times brighter than antibody fluorescence. It is therefore essential to verify that the presence of tracking dye does not compromise the ability to detect other probes being used.</li>
	<li>Failure to obtain a good fit with peak modeling software. Such software allows quantitative comparison of proliferative responses across different populations or stimuli based on precursor frequency or other metrics. Obtaining a good fit, however, requires exclusion of dead/dying cells that can distort dye dilution profiles and matching of the assumptions underlying the model with characteristics of the observed dye dilution profile.</li>
</ol>
Examples given here illustrate how these variables can affect results when using membrane and/or protein dyes to monitor cell proliferation.

Successful use of cell tracking dyes to monitor immune cell function and proliferation involves several critical steps. We describe methods for: 1) obtaining bright, uniform, reproducible label-ing with membrane dyes; 2) selecting fluorochromes and data acquisition conditions; and 3) choosing a model to quantify cell proliferation based on dye dilution.

Fluorescent cell tracking dyes, in combination with flow and image cytometry, are powerful tools with which to study the interactions and fates of ...

Cell Tracking Using Photoconvertible Proteins During Zebrafish Development

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of genetically-encoded fluorescent proteins has proven a valuable strategy for elucidating dynamic morphogenetic processes as they occur in the intact organism. During the past decade, the development of photoactivatable and photoconvertible fluorescent proteins has opened the possibility to investigate the fate of discrete subpopulations of tagged proteins1. Unlike photoactivatable proteins, photoconvertible fluorescent proteins (PCFPs) are readily tracked and imaged in their native emission state prior to photoconversion, making it easier to identify and select regions by optical inspection. PCFPs, such as Kaede2, KikGR3, Dendra4 and EosFP5, can be shifted from green to red upon exposure to UV or blue light due to a His-Tyr-Gly tripeptide sequence which forms a green chromophore that can be photoconverted to a red one by a light-catalyzed &beta;-elimination and subsequent extension of a &pi;-conjugated system3. PCFPs and their monomeric variants are useful tools for tracking cells6-10 and studying protein dynamics11-14, respectively. During recent years, PCFPs have been expressed in different animal model, such as zebrafish6, chicken7,8 and mouse9,10 for cell fate tracking. Here we report a protocol for cell-specific photoconversion of PCFPs in the living zebrafish embryo and further tracking of photoconverted proteins at later developmental stages. This methodology allows studying, in a tissue-specific manner, cell biological events underlying morphogenesis in the zebrafish animal model.

Here, we present a method for the photoactivated switch of photoconvertible fluorescent proteins (PCFPs) in the living zebrafish embryo and further tracking of photoconverted protein at specific time points during development. This methodology allows monitoring of cell biological events underlying different developmental processes in a live vertebrate organism.

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of ...

Techniques for the Analysis of Extracellular Vesicles Using Flow Cytometry

Extracellular Vesicles (EVs) are small, membrane-derived vesicles found in bodily fluids that are highly involved in cell-cell communication and help regulate a diverse range of biological processes. Analysis of EVs using flow cytometry (FCM) has been notoriously difficult due to their small size and lack of discrete populations positive for markers of interest. Methods for EV analysis, while considerably improved over the last decade, are still a work in progress. Unfortunately, there is no one-size-fits-all protocol, and several aspects must be considered when determining the most appropriate method to use. Presented here are several different techniques for processing EVs and two protocols for analyzing EVs using either individual detection or a bead-based approach. The methods described here will assist with eliminating the antibody aggregates commonly found in commercial preparations, increasing signal&#8211;to-noise ratio, and setting gates in a rational fashion that minimizes detection of background fluorescence. The first protocol uses an individual detection method that is especially well suited for analyzing a high volume of clinical samples, while the second protocol uses a bead-based approach to capture and detect smaller EVs and exosomes.

Many different methods exist for the measurement of extracellular vesicles (EVs) using flow cytometry (FCM). Several aspects should be considered when determining the most appropriate method to use. Two protocols for measuring EVs are presented, using either individual detection or a bead-based approach.

Extracellular Vesicles (EVs) are small, membrane-derived vesicles found in bodily fluids that are highly involved in cell-cell communication and help ...

Synchronization of Caulobacter Crescentus for Investigation of the Bacterial Cell Cycle

The cell cycle is important for growth, genome replication, and development in all cells. In bacteria, studies of the cell cycle have focused largely on unsynchronized cells making it difficult to order the temporal events required for cell cycle progression, genome replication, and division. Caulobacter crescentus provides an excellent model system for the bacterial cell cycle whereby cells can be rapidly synchronized in a G0 state by density centrifugation. Cell cycle synchronization experiments have been used to establish the molecular events governing chromosome replication and segregation, to map a genetic regulatory network controlling cell cycle progression, and to identify the establishment of polar signaling complexes required for asymmetric cell division. Here we provide a detailed protocol for the rapid synchronization of Caulobacter NA1000 cells. Synchronization can be performed in a large-scale format for gene expression profiling and western blot assays, as well as a small-scale format for microscopy or FACS assays. The rapid synchronizability and high cell yields of Caulobacter make this organism a powerful model system for studies of the bacterial cell cycle.

Synchronization of Caulobacter Crescentus for Investigation of the Bacterial Cell Cycle

Synchronization of bacterial cells is essential for studies of the bacterial cell cycle and development. Caulobacter crescentus is synchronizable through density centrifugation allowing a rapid and powerful tool for studies of the bacterial cell cycle. Here we provide a detailed protocol for the synchronization of Caulobacter cells.

The cell cycle is important for growth, genome replication, and development in all cells.  In bacteria, studies of the cell cycle have focused largely on ...

LED Thermo Flow &#8212; Combining Optogenetics with Flow Cytometry

Optogenetic tools allow isolated, functional investigations of almost any signaling molecule within complex signaling pathways. A major obstacle is the controlled delivery of light to the cell sample and hence the most popular tools for optogenetic studies are microscopy-based cell analyses and in vitro experiments. The flow cytometer has major advantages over a microscope, including the ability to rapidly measure thousands of cells at single cell resolution. However, it is not yet widely used in optogenetics. Here, we present a device that combines the power of optogenetics and flow cytometry: the LED Thermo Flow. This device illuminates cells at specific wavelengths, light intensities and temperatures during flow cytometric measurements. It can be built at low cost and be used with most common flow cytometers. To demonstrate its utility, we characterized the photoswitching kinetics of Dronpa proteins in vivo and in real time. This protocol can be adapted to almost all optically controlled substances and substantially expands the set of possible experiments. More importantly, it will greatly simplify the discovery and development of new optogenetic tools.

LED Thermo Flow &amp;#8212; Combining Optogenetics with Flow Cytometry

Optically controlled substances are powerful tools to study signaling pathways. To expand the spectrum of possible experiments, we developed a device for studying optically controlled substances in real time using flow cytometry: the LED Thermo Flow.

Optogenetic tools allow isolated, functional investigations of almost any signaling molecule within complex signaling pathways. A major obstacle is the ...

Determination of the Relative Cell Surface and Total Expression of Recombinant Ion Channels Using Flow Cytometry

Inherited or de novo mutations in cation-selective channels may lead to sudden cardiac death. Alteration in the plasma membrane trafficking of these multi-spanning transmembrane proteins, with or without change in channel gating, is often postulated to contribute significantly in this process. It has thus become critical to develop a method to quantify the change of the relative cell surface expression of cardiac ion channels on a large scale. Herein, a detailed protocol is provided to determine the relative total and cell surface expression of cardiac L-type calcium channels CaV1.2 and membrane-associated subunits in tsA-201 cells using two-color fluorescent cytometry assays. Compared with other microscopy-based or immunoblotting-based qualitative methods, flow cytometry experiments are fast, reproducible, and large-volume assays that deliver quantifiable end-points on large samples of live cells (ranging from 104 to 106 cells) with similar cellular characteristics in a single flow. Constructs were designed to constitutively express mCherry at the intracellular C-terminus (thus allowing a rapid assessment of the total protein expression) and express an extracellular-facing hemagglutinin (HA) epitope to estimate the cell surface expression of membrane proteins using an anti-HA fluorescence conjugated antibody. To avoid false negative, experiments were also conducted in permeabilized cells to confirm the accessibility and proper expression of the HA epitope. The detailed procedure provides: (1) design of tagged DNA (deoxyribonucleic acid) constructs, (2) lipid-mediated transfection of constructs in tsA-201 cells, (3) culture, harvest, and staining of non-permeabilized and permeabilized cells, and (4) acquisition and analysis of fluorescent signals. Additionally, the basic principles of flow cytometry are explained and the experimental design, including the choice of fluorophores, titration of the HA antibody and control experiments, is thoroughly discussed. This specific approach offers objective relative quantification of the total and cell surface expression of ion channels that can be extended to study ion pumps and plasma membrane transporters.

Inherited cardiac arrhythmias are often caused by mutations that alter the surface delivery of one or more ion channels. Here, we adapt flow cytometry assays to provide a quantification of the relative total and cell surface protein expression of recombinant ion channels expressed in tsA-201 cells.

Inherited or de novo mutations in cation-selective channels may lead to sudden cardiac death. Alteration in the plasma membrane trafficking of these ...

Analysis of Cell Suspensions Isolated from Solid Tissues by Spectral Flow Cytometry

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to the analysis of a few fluorochromes, currently there are dozens of different fluorescent dyes, and up to 14-18 different dyes can be combined at a time. However, several limitations still impair the analytical capabilities. Because of the multiplicity of fluorescent probes, data analysis has become increasingly complex due to the need of large, multi-parametric compensation matrices. Moreover, mutant mouse models carrying fluorescent proteins to detect and trace specific cell types in different tissues have become available, so the analysis (by flow cytometry) of auto-fluorescent cell suspensions obtained from solid organs is required. Spectral flow cytometry, which distinguishes the shapes of emission spectra along a wide range of continuous wavelengths, addresses some of these problems. The data is analyzed with an algorithm that replaces compensation matrices and treats auto-fluorescence as an independent parameter. Thus, spectral flow cytometry should be capable of discriminating fluorochromes with similar emission peaks and can provide a multi-parametric analysis without compensation requirements.
This protocol describes the spectral flow cytometry analysis, allowing for a 21-parameter (19 fluorescent probes) characterization and the management of an auto-fluorescent signal, providing high resolution in minor population detection. The results presented here show that spectral flow cytometry presents advantages in the analysis of cell populations from tissues difficult to characterize in conventional flow cytometry, such as the heart and the intestine. Spectral flow cytometry thus demonstrates the multi-parametric analytical capacity of high-performing conventional flow cytometry without the requirement for compensation and enables auto-fluorescence management.

This article describes spectral cytometry, a new approach in flow cytometry that uses the shapes of emission spectra to distinguish fluorochromes. An algorithm replaces compensations and can treat auto-fluorescence as an independent parameter. This new approach allows for the proper analysis of cells isolated from solid organs.

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to ...

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

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

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

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