We thank Andy Johnson of the Biomedical Research Center at UBC for assistance with FACS analysis. Cancer Research Funding in the Wong Laboratory is provided by the Canadian Cancer Society Research Institute (operating grant #019250) and from Research Reinvestment funds of the Faculty of Pharmaceutical Sciences, UBC. JMYW is supported by the Canada Research Chairs and the Michael Smith Foundation for Health Research Career Development programs.

1. Cell Preparation
<img src="/files/ftp_upload/4045/4045fig0.jpg" alt="Figure 0" /> 
<ol>
 <li> Plate cells to achieve a density of approximately 60% confluency. Cells must be in log phase at the time of collection. For mcf7 cells, this is accomplished by seeding at 5 x 105 cells / 10 cm plate in the appropriate media. HT29 and LS180 cells are seeded at 6 x 105 cells / 6 cm plate. We used DMEM media supplemented with 10% FBS and 1x penicillin/streptomycin. Be sure to seed plates for the correct positive and negative controls in addition to your test samples. They include the following: </li>
</ol>
 <table border="1">
 <tbody>
 <tr>
 <td>i</td> 
 <td>positive control</td>
 <td>BrdU only</td>
 </tr>
 <tr>
 <td>ii</td>
 <td>positive control</td>
 <td>PI only</td>
 </tr>
 <tr>
 <td>iii</td>
 <td>negative control</td>
 <td>BrdU negative, PI negative</td>
 </tr>
 </tbody>
 </table>
(It should be noted that a preliminary experiment with multiple timepoints such as the one outlined in this protocol can be used to narrow down the time frame during which to carry out future collections.) 
<ol start="2">
 <li> The plated cells are incubated at 37 &deg;C, 5% CO2 for 24-48 h, thereby allowing cells to recover and attach. </li>
 <li> To pulse label cells with bromodeoxyuridine (BrdU), DMEM is replaced with fresh media containing 10 &mu;M BrdU. Cells are incubated for 1 h at 37 &deg;C, 5% CO2 to allow for BrdU incorporation into the DNA. Be sure to leave one plate untreated to act as the negative control. </li>
 <li> The pulse-labeling media is removed and the cells are rinsed briefly with 1X PBS.</li>
 <li> Fresh growth media is added (minus the BrdU) and cells are allowed to continue to incubate at 37 &deg;C, 5% CO2 until the appropriate timepoint for harvesting is reached. </li>
</ol>
2. Harvesting and Fixation
<ol>
 <li> Untreated cells from step 1.3 are considered the zero timepoint. This sample can be harvested together with the 1 h timepoint which is collected immediately after BrdU treatment. </li>
 <li> To harvest the cells, media is removed and the plates are rinsed once with 1X PBS. </li>
 <li> Cells are trypsinized, collected in culturing media and subsequently pelleted by centrifugation at 1500 rpm for 5 min Supernatant is discarded. </li>
 <li> Rinse 1-10 x 106 cells in 5 ml ice cold 1X PBS. Centrifuge at 1500 rpm for 5 min Supernatant is discarded. </li>
 <li> The cell pellet is then resuspended in 100 &mu;l of ice-cold PBS/1% FBS. Addition of 1% FBS in PBS aids in preventing cell clumping. </li>
 <li> Add these cells drop by drop to 5 ml of -20 &deg;C, 70% ethanol to fix the cells. </li>
 <li> Incubate on ice for 30 min or store at 4 &deg;C overnight. This is an ideal step at which to stop the collection of samples from the various timepoints. The samples can be left in ethanol for several days, allowing them to be processed simultaneously through the remainder of the protocol. </li>
</ol>
3. BrdU and PI Staining
<ol>
 <li> Pellet cells by centrifugation for 5 min at 1500 rpm.</li>
 <li> Remove the fixative, but leave ~50 &mu;l in which to loosen the pellet by vortexing. </li>
 <li> To denature the DNA, slowly add 1 ml of 2N HCL/Triton X-100 dropwise while vortexing. Incubate samples at room temperature for 30 min. </li>
 <li> Pellet cells by centrifuging samples for 5 min at 1500 rpm. Aspirate and discard the supernatant. </li>
 <li> Resuspend the cell pellet in 1 ml of 0.1M sodium tetraborate, pH 8.5, to neutralize the denaturation step. </li>
 <li> Centrifuge cells for 5 min at 1500 rpm. Aspirate and discard the supernatant. </li>
 <li> Resuspend the cell pellet in 75 &mu;l of BrdU staining mix (50 &mu;l 0.5% tween 20/1% BSA/PBS + 20 &mu;l FITC conjugated anti-BrdU + 5 &mu;l 10 mg/ml RNase). </li>
 <li> Incubate at room temperature for 45 min protected from light. </li>
 <li> To pellet cells, centrifuge the samples for 5 min at 1500 rpm. Aspirate and discard the supernatant. </li>
 <li> Resuspend the cell pellet in 1 ml of PBS containing 5 &mu;g/ml propidium iodide. </li>
</ol>
4. Flow Cytometry
<ol>
 <li> To analyze the cells, a flow cytometer equipped with a 488 nm laser and the appropriate filters is necessary. Analysis software such as CellQuest is necessary to create the plots outlined below. </li>
 <li> When running the sample through the flow cytometer, create a forward scatter (SSC-H) vs. side scatter (FSC-H) plot to ensure the proper size distribution of cells (Figure 1A). Both of these parameters are plotted on a linear scale. </li>
 <li> Simultaneously view a plot of FL3-H (PI stain) vs. FL2-W (Figure 1B). This plot is used to create a gate (R3) to isolate the fraction of cells that are within the normal distribution pattern of the cell cycle. In general, this distribution pattern is characterized by two clusters of cells from the 2N and 4N DNA content of PI staining representing the G1 and G2 phases of the cell cycle, respectively. A string of cells located between these 2 clusters is representative of ongoing DNA replication that occurs in the S phase of the cell cycle. </li>
 <li> Create a plot displaying the gated cells (R3) from step 4.3, with FL1-H (FITC, log plot) on the y-axis and PI (linear plot) on the x-axis (Figure 1C). This plot will be used to set the remaining parameters using the various controls outlined in 1.1, as well as to collect data for analysis. </li>
 <li> Place the PI only stained sample onto the cytometer. Adjust the gain to place G1 at ~200 on the x-axis. This will be easier to visualize in a histogram plot of the PI stain. </li>
 <li> A second control involves running a BrdU-only sample on the flow cytometer. Adjust the gain so that the 2 populations of cells (BrdU positive vs. BrdU negative) appear on the plot. Ideally, the BrdU negative cells are positioned to appear just below 10-1. </li>
 <li> The final control is the BrdU/PI negative sample. Run this negative control to ensure that cells do not appear in any of the upper or right hand quadrants. </li>
 <li> Once the parameters of the various plots have been set, the flow cytometer is calibrated and ready for processing of BrdU and PI stained cells. A minimum of 10,000 cells that are gated in the correct PI fraction (refer to step 4.3) must be read per collection. </li>
 <li> To analyze the collected data, software such as FlowJo or FacsDiva are utilized. Each of the above mentioned plots can be recreated in these programs and quantitative and statistical analysis performed. </li>
 <li> The endpoint of the analysis involves several successive steps. Creating a plot of FITC vs. PI with cells gated as PI positive from the PI vs. FL2-W plot allows one to distinguish the cycling population. A second gate is created from this plot by isolating the BrdU positive population. Expressing this distinctive population on a histogram plot with PI on the x-axis allows one to track the time course of cell cycle progression. This can be further visualized by plotting the number of BrdU-positive cells with G1 or G2 content as a function of time. </li>
</ol>
5. Representative Results
Normally cycling cells stained with propidium iodide have distinct peaks at G1 and G2, corresponding to cells containing 2N and 4N DNA content, respectively (Figure 2). Pulse labeling with BrdU allows for selective labeling of a sub-population of cells that are actively synthesizing DNA (i.e. S phase). Shortly after removal of the BrdU reagent, all labeled cells are in S phase (Figure 3). By restricting labeling to a short pulse one is able to follow this now distinct sub-population of cells throughout numerous time points as they pass through the subsequent phases of the cell cycle. This can be visualized at the 1 h time point in Figure 3 as a distinct lack of G1 and G2 peaks. 
Additional information can be derived from the BrdU labeling step. Not only can the proportion of actively dividing cells be measured, but an estimate of cell cycle stage distribution between two samples can be determined as well. By collecting cells at equally spaced intervals following the removal of the BrdU reagent, cells can be traced as they continue to cycle, progress to G2 and ultimately move through to cell division of the original cells, finally emerging as BrdU-positive daughter cells with G1 DNA content (Figure 4). An example of cell cycle phase quantification and kinetic analysis is provided in Figure 5.
<img src="/files/ftp_upload/4045/4045fig1.jpg" alt="Figure 1" /> 
Figure 1. (A) Forward Scatter versus Side Scatter Plot of a representative mcf7 cell population. (B) PI versus Width (FL-W) Plot of a representative mcf7 cell population. Gating is shown to exclude cell doublets in the final analysis (R3). Cell doublets will have greater pulse width than a single cell, as they take longer to pass through the laser beam and therefore can be excluded from the analysis. (C) PI versus FITC (BrdU) Plot of a representative mcf7 cell population. Gating is shown to include only the FITC (BrdU) positive cells (R4).
<img src="/files/ftp_upload/4045/4045fig2.jpg" alt="Figure 2" /> 
Figure 2. Representative histogram plot of a total population of normally cycling cells.
<img src="/files/ftp_upload/4045/4045fig3.jpg" alt="Figure 3" /> 
Figure 3. Histogram plot of cells that have been collected 1h after the removal of the BrdU pulse, after gating for FITC (BrdU)-positive cells. BrdU-positive cells at an early timepoint following the removal of label show PI profiles that correspond to samples displaying DNA content consistent with cells that are in the S-phase of the cell cycle, confirming the successful labeling of cells only during DNA synthesis.
<img src="/files/ftp_upload/4045/4045fig4.jpg" alt="Figure 4" /> 
Figure 4. Comparison of cell cycle progression kinetics between cancer cell lines, colorectal cancers HT29 (A) and LS180 (B), as well as breast cancer mcf7 (C). Cells were collected every hour for 8 hours, following the removal of the BrdU pulse. In this experiment, we observed a clear profile of accelerated cell cycle progression through the G2 phase of the cell cycle in the colorectal cell line LS180. Comparing the kinetics between the two profiles of colorectal cancer cells, the emergence of a G1 peak is evident at T= 4h post-BrdU in the LS180 cells, compared with the corresponding time point for the HT-29 cell line which lacks this peak. Compared with either one of the two colorectal cell lines, mcf7 cells are cycling at a significantly reduced rate. <a href="https://www.jove.com/files/ftp_upload/4045/4045fig4large.jpg">Click here to view larger figure</a>.
<img src="/files/ftp_upload/4045/4045fig5.jpg" alt="Figure 5" /> 
Figure 5. (A) Quantitative cell cycle phase analysis of BrdU-labeled cancer cells. The Dean/Jett/Fox algorithm was applied to HT29, LS180 and mcf7 (illustrated in green). The resulting cell cycle phase distributions from each sample are expressed as % total BrdU-positive cells for each phase. Only select time points for each cell line are displayed, as the analyses for some of the earlier time points produced invalid results. In these earlier time points, all BrdU-positive cells are in S-phase and therefore lack distinct G1 and G2 peaks, which are necessary for algorithm application. (B) Histograms of cancer cells showing the kinetics of progression through G2/M phases of the cell cycle. Progression through G2/M phases is fastest for LS180 cells, followed by HT29 and mcf7. <a href="https://www.jove.com/files/ftp_upload/4045/4045fig5large.jpg">Click here to view larger figure</a>.
<img src="/files/ftp_upload/4045/4045figS1.jpg" alt="Figure S1" /> 
Figure S1. Flow cytometry controls. mcf7 cells are shown as (A) negative control where cells are neither PI nor BrdU stained. (B) PI only stained. (C) BrdU-FITC labeled samples. 
<img src="/files/ftp_upload/4045/4045figS2.jpg" alt="Figure S2" /> 
Figure S2. Schematic illustration showing the relationship between fluorescent emission spectrum of PI versus FITC (BrdU). The spectral collection windows for fluorescent channel 1 (FL1, for FITC) and fluorescent channel 3 (FL3, for PI) are shown in corresponding boxes. There is no fluorophore spectra overlap between FL1 and FL3 detection. Evidently, fluorescence compensation is not necessary when experimental data from PI and FITC-BrdU co-labeled cells are collected in the FL1/FL3 channels respectively. Fluorescence spectrums were obtained from BD BioSciences website: <a href="http://www.bdbiosciences.com/research/multicolor/spectrum_viewer" target="_blank">http://www.bdbiosciences.com/research/multicolor/spectrum_viewer</a>. <a href="https://www.jove.com/files/ftp_upload/4045/4045figS2large.jpg" target="_blank">Click here to view larger figure</a>.

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	<li>Musgrove, E. A., Caldon, C. E., Barraclough, J., Stone, A., Sutherland, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cyclin+D+as+a+therapeutic+target+in+cancer.">Cyclin D as a therapeutic target in cancer.</a> Nat. Rev. Cancer. 11, 558-572 (2011).</li><li>Molchadsky, A., Rivlin, N., Brosh, R., Rotter, V., Sarig, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=p53+is+balancing+development,+differentiation+and+de-differentiation+to+assure+cancer+prevention.">p53 is balancing development, differentiation and de-differentiation to assure cancer prevention.</a> Carcinogenesis. 31, 1501-1508 (2010).</li><li>Dickson, M. A., Schwartz, G. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+cell-cycle+inhibitors+for+cancer+therapy.">Development of cell-cycle inhibitors for cancer therapy.</a> Curr. Oncol. 16, 36-43 (2009).</li><li>Banfalvi, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+cycle+synchronization+of+animal+cells+and+nuclei+by+centrifugal+elutriation.">Cell cycle synchronization of animal cells and nuclei by centrifugal elutriation.</a> Nat. 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We have nothing to disclose.

By combining flow cytometry with BrdU incorporation, we have the necessary tools to study cell cycle kinetics. The distinctive property of BrdU to function as a thymidine analogue is what allows for DNA content quantification of a cycling cell. The incorporation of BrdU into a growing daughter DNA strand during the synthesis phase of the cell cycle is what allows one to follow a subpopulation of cycling cells through the replication of DNA in S phase, to a period of growth at G2 and ultimately to cell division. Daught...

<table border="1">
<tbody>
 <tr>
 <td>Reagent</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>bromodeoxyuridine</td>
 <td>Becton Dickinson</td>
 <td>55089</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>propidium iodide</td>
 <td>Sigma</td>
 <td>287075</td>
 <td>1mg/ml stock</td>
 </tr>
 <tr>
 <td>FITC anti-BrdU</td>
 <td>Becton Dickinson</td>
 <td>347583</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>sodium tetraborate</td>
 <td>Fisher</td>
 <td>S80172</td>
 <td>0.1M, pH 8.5</td>
 </tr>
 <tr>
 <td>FACS Caliber</td>
 <td>Becton Dickinson</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry at JoVE.com

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

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 ...

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21.0K Views.  University of British Columbia. 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.

Video: Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

Measuring Exocytosis in Neurons Using FM Labeling

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time imaging of vesicles labeled with FM dye to monitor the rate of presynaptic vesicle release.  FM4-64 is a red fluorescent amphiphilic styryl dye that embeds into the membranes of synaptic vesicles as endocytosis is stimulated.  Lipophilic interactions cause the dye to greatly increase in fluorescence, thus emitting a bright signal when associated with vesicles and a nominal one when in the extracellular fluid. After a wash step is used to help remove external dye within the plasma membrane, the remaining FM is concentrated within the vesicles and is then expelled when exocytosis is induced by another round of electrical stimulation. The rate of vesicles release is measured from the resulting decrease in fluorescence.  Since FM dye can be applied external and transiently, it is a useful tool for determining rates of exocytosis in neuronal cultures, especially when comparing the rates between transfected synapses and neighboring control boutons.



The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission. Here we used real-time imaging of vesicles labeled with the red fluorescent dye FM 4-64 to measure the rate of presynaptic vesicle release in hippocampal neuronal cultures.

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time ...

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 ...

Flow Cytometry-based Purification of S. cerevisiae Zygotes

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae zygotes result from the fusion of haploid cells of distinct mating type (MATa, MATalpha) and give rise to corresponding stable diploids that successively generate as many as 20 diploid progeny as a result of their strikingly asymmetric mitotic divisions 2. Zygote formation is orchestrated by a complex sequence of events: In this process, soluble mating factors bind to cognate receptors, triggering receptor-mediated signaling cascades that facilitate interruption of the cell cycle and culminate in cell-cell fusion. Zygotes may be considered a model for progenitor or stem cell function.
Although much has been learned about the formation of zygotes and although zygotes have been used to investigate cell-molecular questions of general significance, almost all studies have made use of mating mixtures in which zygotes are intermixed with a majority population of haploid cells 3-8. Many aspects of the biochemistry of zygote formation and the continuing life of the zygote therefore remain uninvestigated.
Reports of purification of yeast zygotes describe protocols based on their sedimentation properties 9; however, this sedimentation-based procedure did not yield nearly 90% purity in our hands. Moreover, it has the disadvantage that cells are exposed to hypertonic sorbitol. We therefore have developed a versatile purification procedure. For this purpose, pairs of haploid cells expressing red or green fluorescent proteins were co-incubated to allow zygote formation, harvested at various times, and the resulting zygotes were purified using a flow cytometry-based sorting protocol. This technique provides a convenient visual assessment of purity and maturation. The average purity of the fraction is approximately 90%. According to the timing of harvest, zygotes of varying degrees of maturity can be recovered. The purified samples provide a convenient point of departure for "-omic" studies, for recovery of initial progeny, and for systematic investigation of this progenitor cell.

Flow Cytometry-based Purification of S. cerevisiae Zygotes

To purify zygotes of S. cerevisiae, haploid cells of opposite mating type were engineered to express red or green fluorescent proteins, co-incubated to allow zygote formation, and fractionated using a flow cytometry-based protocol. The highly-enriched fraction enables subsequent "-omic" studies, recovery of initial progeny, and systematic investigation of zygote morphogenesis.

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae ...

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 ...

Measuring Intracellular Ca2+ Changes in Human Sperm using Four Techniques: Conventional Fluorometry, Stopped Flow Fluorometry, Flow Cytometry and Single Cell Imaging

Spermatozoa are male reproductive cells especially designed to reach, recognize and fuse with the egg. To perform these tasks, sperm cells must be prepared to face a constantly changing environment and to overcome several physical barriers. Being in essence transcriptionally and translationally silent, these motile cells rely profoundly on diverse signaling mechanisms to orient themselves and swim in a directed fashion, and to contend with challenging environmental conditions during their journey to find the egg. In particular, Ca2+-mediated signaling is pivotal for several sperm functions: activation of motility, capacitation (a complex process that prepares sperm for the acrosome reaction) and the acrosome reaction (an exocytotic event that allows sperm-egg fusion). The use of fluorescent dyes to track intracellular fluctuations of this ion is of remarkable importance due to their ease of application, sensitivity, and versatility of detection. Using one single dye-loading protocol we utilize four different fluorometric techniques to monitor sperm Ca2+ dynamics. Each technique provides distinct information that enables spatial and/or temporal resolution, generating data both at single cell and cell population levels.

Measuring Intracellular Ca2+ Changes in Human Sperm using Four Techniques: Conventional Fluorometry, Stopped Flow Fluorometry, Flow Cytometry and Single Cell Imaging

Intracellular Ca2+ dynamics are very important in sperm physiology and Ca2+-sensitive fluorescent dyes constitute a versatile tool to study them. Population experiments (fluorometry and stopped flow fluorometry) and single cell experiments (flow cytometry and single cell imaging) are used to track spatio-temporal [Ca2+] changes in human sperm cells.

Spermatozoa  are male reproductive cells especially designed to reach, recognize and fuse  with the egg. To perform these tasks, sperm cells must be ...

Metabolic Labeling of Leucine Rich Repeat Kinases 1 and 2 with Radioactive Phosphate

Leucine rich repeat kinases 1 and 2 (LRRK1 and LRRK2) are paralogs which share a similar domain organization, including a serine-threonine kinase domain, a Ras of complex proteins domain (ROC), a C-terminal of ROC domain (COR), and leucine-rich and ankyrin-like repeats at the N-terminus. The precise cellular roles of LRRK1 and LRRK2 have yet to be elucidated, however LRRK1 has been implicated in tyrosine kinase receptor signaling1,2, while LRRK2 is implicated in the pathogenesis of Parkinson's disease3,4. In this report, we present a protocol to label the LRRK1 and LRRK2 proteins in cells with 32P orthophosphate, thereby providing a means to measure the overall phosphorylation levels of these 2 proteins in cells. In brief, affinity tagged LRRK proteins are expressed in HEK293T cells which are exposed to medium containing 32P-orthophosphate. The 32P-orthophosphate is assimilated by the cells after only a few hours of incubation and all molecules in the cell containing phosphates are thereby radioactively labeled. Via the affinity tag (3xflag) the LRRK proteins are isolated from other cellular components by immunoprecipitation. Immunoprecipitates are then separated via SDS-PAGE, blotted to PVDF membranes and analysis of the incorporated phosphates is performed by autoradiography (32P signal) and western detection (protein signal) of the proteins on the blots. The protocol can readily be adapted to monitor phosphorylation of any other protein that can be expressed in cells and isolated by immunoprecipitation.

Leucine rich repeat kinases 1 and 2 (LRRK1 and LRRK2) are multidomain proteins which encode both GTPase and kinase domains and which are phosphorylated in cells. Here, we present a protocol to label LRRK1 and LRRK2 in cells with 32P orthophosphate, thereby providing a means to measure their overall cellular phophorylation levels.

Leucine rich repeat kinases 1 and 2 (LRRK1 and LRRK2) are paralogs which share a similar domain organization, including a serine-threonine kinase domain, ...

Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures

Plasma membrane microdomains are features based on the physical properties of the lipid and sterol environment and have particular roles in signaling processes. Extracting sterol-enriched membrane microdomains from plant cells for proteomic analysis is a difficult task mainly due to multiple preparation steps and sources for contaminations from other cellular compartments. The plasma membrane constitutes only about 5-20% of all the membranes in a plant cell, and therefore isolation of highly purified plasma membrane fraction is challenging. A frequently used method involves aqueous two-phase partitioning in polyethylene glycol and dextran, which yields plasma membrane vesicles with a purity of 95% 1. Sterol-rich membrane microdomains within the plasma membrane are insoluble upon treatment with cold nonionic detergents at alkaline pH. This detergent-resistant membrane fraction can be separated from the bulk plasma membrane by ultracentrifugation in a sucrose gradient 2. Subsequently, proteins can be extracted from the low density band of the sucrose gradient by methanol/chloroform precipitation. Extracted protein will then be trypsin digested, desalted and finally analyzed by LC-MS/MS. Our extraction protocol for sterol-rich microdomains is optimized for the preparation of clean detergent-resistant membrane fractions from Arabidopsis thaliana cell cultures.
We use full metabolic labeling of Arabidopsis thaliana suspension cell cultures with K15NO3 as the only nitrogen source for quantitative comparative proteomic studies following biological treatment of interest 3. By mixing equal ratios of labeled and unlabeled cell cultures for joint protein extraction the influence of preparation steps on final quantitative result is kept at a minimum. Also loss of material during extraction will affect both control and treatment samples in the same way, and therefore the ratio of light and heave peptide will remain constant. In the proposed method either labeled or unlabeled cell culture undergoes a biological treatment, while the other serves as control 4.

Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures

Here we describe a robust method for the fractionation of plant plasma membranes into detergent resistant and detergent soluble membranes based on a mixture of unlabeled and in vivo fully 15N labeled Arabidopsis thaliana cell cultures. The procedure is applied for comparative proteomic studies to understand signaling processes.

Plasma membrane microdomains are features based on the physical properties of the lipid and sterol environment and have particular roles in signaling ...

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 ...

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 ...

Measuring Endoreduplication by Flow Cytometry of Isolated Tuber Protoplasts

Endoreduplication, the replication of a cell's nuclear genome without subsequent cytokinesis, yields cells with increased DNA content and is associated with specialization, development and increase in cellular size. In plants, endoreduplication seems to facilitate the growth and expansion of certain tissues and organs. Among them is the tuber of potato (Solanum tuberosum), which undergoes considerable cellular expansion in fulfilling its function of carbohydrate storage. Thus, endoreduplication may play an important role in how tubers are able to accommodate this abundance of carbon. However, the cellular debris resulting from crude nuclear isolation methods of tubers, methods that can be used effectively with leaves, precludes the estimation of the tuber endoreduplication index (EI). This article presents a technique for assessing tuber endoreduplication through the isolation of protoplasts while demonstrating representative results obtained from different genotypes and compartmentalized tuber tissues. The major limitations of the protocol are the time and reagent costs required for sample preparation as well as relatively short lifespan of samples after lysis of protoplasts. While the protocol is sensitive to technical variation, it represents an improvement over traditional methods of nuclear isolation from these large specialized cells. Possibilities for improvements to the protocol such as recycling enzyme, the use of fixatives, and other alterations are proposed.

The protocol described herein is a method for measuring endoreduplication within tubers of potato (Solanum tuberosum). It includes plasmolysis and protoplast extraction steps to decrease the noise and debris in downstream flow cytometric analysis.

Endoreduplication, the replication of a cell's nuclear genome without subsequent cytokinesis, yields cells with increased DNA content and is associated ...