The authors wish to thank the Canadian Institutes of Health Research (CIHR) and the Canadian Cancer Society for funding cell cycle research in their laboratory. MJC is a recipient of an MD/PhD fellowship from the CIHR. MJC and MA are members of the CaRTT training program.

1. Labeling and fixing of cells
<ol>
	<li>Add 1 &mu;L of Cell Proliferation Labeling Reagent (BrdU) per mL of cell culture medium (1 to 1000 dilution) 1 hour before harvesting. The labeling period may need to be lengthened for slower growing cells.</li>
	<li>To harvest cells, aspirate culture medium and wash thoroughly with phosphate buffered saline (PBS). Repeat to thoroughly remove traces of medium.</li>
	<li>Wash cultures quickly a third time with PBS containing 3mM EDTA and aspirate thoroughly.</li>
	<li>Add a small volume of PBS containing 3mM EDTA to each dish, 0.5 mL for a 6 cm dish is ideal. Incubate at 22&deg;C for approximately 5 minutes to detach cells. Transfer to a 15 mL conical tube.</li>
	<li>Centrifuge cells at 500 x g for 5 minutes to pellet, remove supernatant, and resuspend thoroughly in 100 &mu;L of PBS.</li>
	<li>Fix cells by adding 5 mL of 95% EtOH, dropwise, while vortexing. At this step, cells can be stored at 4&deg;C for at least a month.</li>
</ol>
2. Denaturing and staining of BrdU and DNA
<ol>
	<li>Centrifuge cells at 500 x g for 5 minutes to pellet cells and remove 95% EtOH. Resuspend in 1 mL of 2N HCl and 0.5% Tx-100 by adding in a drop wise fashion while vortexing. Incubate at room temperature for 30 minutes.</li>
	<li>Centrifuge as before in section 2.1 and carefully aspirate supernatant since the cells form a very loose pellet at this step. Gently resuspend in 1 mL of 0.1M NaB4O7 (pH 8.5) and incubated for at least 30 minutes at room temperature.</li>
	<li>Pellet cells as above in section 2.1 and resuspend in 0.5 mL of antibody solution (PBS containing 1% BSA and 0.2% Tween-20) with mouse anti-BrdU antibodies diluted 1 to 50. Incubate at room temperature for 30 minutes.</li>
	<li>Pellet cells again and resuspend in 50 mL of antibody solution containing rabbit anti-mouse secondary antibodies conjugated to FITC diluted 1 to 25. Incubate for 30 minutes on benchtop and protect from light.</li>
	<li>Centrifuge cells a final time and resuspend in 0.5 mL of propidium iodide and RNase solution (PI-RNAse solution, PBS with 1% BSA, 10 mg/mL propidium iodide, 0.25 mg/mL RNase A) and incubate in the dark at 37&deg;C for 30 minutes. Alternatively RNA can be digested overnight at 4&deg;C in the dark.</li>
	<li>Pass solution through a cell strainer to remove aggregates and collect single cells in an appropriate tube for sample uptake on your flow cytometer. Again, samples should be protected from direct exposure to light.</li>
</ol>
3. Analysis by flow cytometry
<ol>
	<li>Cells should be analyzed by a standard flow cytometer that is capable of discriminating against doublets (two cells that pass through the flow cell as one) with appropriate detection capability for propidium iodide and FITC. Single cells can be detected in this flow cytometry application by gating for events based on forward scatter and side scatter populations, and by using the properties of DNA staining by propidium iodide. The appearance of a doublet discriminating dot plot varies between flow cytometers and relies on propidium iodide staining properties, see your local flow cytometry facility for advice on setting up a doublet discriminator in your analysis. In general, in an asynchronously proliferating population of cells, single cells are usually the two most abundant peaks (2N and 4N DNA) in the doublet discriminating plot and a gate should be drawn around them. This will select single cell events that are used for all subsequent analysis below.</li>
	<li>The first sample to be analyzed should be an asynchronously proliferating culture that serves as a positive control for staining and flow cytometer set-up. Adjust the sensitivity of photomultiplier tubes for propidium iodide staining such that the 2N and 4N peaks from singlet cells are centered at 200 and 400 (arbitrary units) on the X-axis. We often stain an abundant supply of these cells to ensure that all parameters of the flow cytometer have been adjusted appropriately without using up this sample. This positive control is also helpful for new users to become more familiar with the operation of the flow cytometer.</li>
	<li>Adjust the sensitivity of the photomultiplier tube for FITC detection such that G1 and G2/M populations are above background. BrdU positive cells should be approximately 10 times brighter and the Y-axis should be displayed as a logarithmic scale. It is sometimes helpful to include a negative control for BrdU staining (by replacing the anti-BrdU antibodies with non-specific IgG in step 2.3 above) to clearly distinguish positively stained cells.</li>
	<li>Adjust compensation between propidium iodide and FITC such that single events captured by the flow cytometer form a horse shoe shaped arc from G1 in the lower left up to S-phase and down into the lower right for G2/M. Please see the following reference for a more in depth discussion of the principles and use of flow cytometry and references there in for specific applications2.</li>
</ol>
4. Data analysis
<ol>
	<li>5 000 to 10 000 single cell events with the desired range of DNA content should be collected for each sample in order to have confidence that the population of cells in the culture have been thoroughly sampled.</li>
	<li>Once cells have been measured for propidium iodide and BrdU content they need to be assigned to the G1, S, or G2/M phases. Do this by drawing gates around the two BrdU negative populations centered at 200 and 400 (G1 and G2/M respectively). Everything above these boxes should be included in a single gate that measures S-phase (Fig. 1A). The percentage of cells in each gate represents the relative number of cells in G1, S, and G2/M (Fig. 1B).</li>
</ol>
5. Alternate protocol for analysis of cell cycle in a mixed population of cells
<ol>
	<li>This alternate protocol is most helpful when transfection of expression vectors routinely results in a low percentage of cells expressing the gene product of interest, or when drug treatment to select a pure population of cells is impractical. This results in a relatively small proportion of cells of interest being contaminated with a large population of untransduced cells. In this case, co-transfect plasmids expressing your gene or shRNA of interest along with a plasmid that expresses a marker for detecting transfected cells by flow cytometry, such as a membrane anchored GFP3, or a foreign cell surface marker such as CD194 or CD205.</li>
	<li>For the purposes of this protocol we will use CD20 staining as an example, however, staining for CD19 is essentially identical. In the case of transfection with CD20, there is no need to BrdU label, simply harvest cells as described in steps 1.2 to 1.5 above and stain by adding 20 &mu;L FITC conjugated anti-CD20 antibodies to the cells on ice for 20 minutes in the dark. Fix cells as described in step 1.6. Cells can again be stored for at least a month at 4&deg;C in the dark. For detection of GFP, omit the antibody staining from this step and fix and store cells as described.</li>
	<li>Re-hydrate the cells by pelleting in the centrifuge at 500 x g for 5 minutes and resuspending in 5 mL of PBS containing 1% BSA.</li>
	<li>Re-pellet the cells at 500 x g for 5 minutes and resuspend cells in 0.5 mL of propidium iodide and RNase solution as described above in section 2.5.</li>
	<li>Continue to follow cell preparation instructions in 2.6 and analysis instructions in 3.1 and 3.2.</li>
	<li>Adjust the sensitivity of the photomultiplier tube for FITC detection such that unstained cells are above background. Ideally, CD20-FITC positive cells will be 10X to 100X more intensely stained than background and the Y-axis of this plot should be displayed as a logarithmic scale (Fig. 2A). CD20 positive cells that are 10X brighter than background (and with propidium iodide staining between 200 and 400) should be selected for display in a propidium iodide versus cell counts histogram as shown in Fig. 2C. For cell lines that express markers that are readily stained brightly (ie. 10X to 100X above background) the CD20 positive cut-off should be set at 10X background. Inclusion of weaker staining cells increases variability in these experiments, presumably because the gene of interest is also weakly expressed in these cells. For cell lines that only yield weakly stained CD20 positives (ie. less than 10X background), determination of a cut off should be made using a negative control consisting of CD20 stained, untransfected cells.</li>
	<li>At least 1000 CD20 positive events should be collected for each sample. Experiments with fewer that 1000 events will produce high variability. Software packages such as ModFit LT (Verity Software), Multi Cycle AV (Phoenix Flow Systems), and Flow Jo (Tree Star) use mathematical estimates of the G1, S, and G2/M populations that contribute to the shape of the curve in histograms such as those shown in Fig. 2B and C.</li>
</ol>
6. Representative Results:
We provide three examples of experimental cell cycle analyses using our approaches. The first uses retroviral expression of the cyclin dependent kinase inhibitor p27KIP1 in mouse embryonic fibroblasts. Twenty four hours after drug selection for viral infection was complete, cells were pulse labeled with BrdU for one hour. In this experiment ectopic expression of the inhibitor is used to arrest proliferation of the cells (Fig. 3A and B). As shown in Fig. 3B little BrdU positive events are evident in the S-phase gate in response to p27. Likewise, the percentage of cells in S-phase for p27 expressing cells is quite low as diagramed in Fig. 3C. This type of analysis has been very effective in characterizing the cell cycle control defects in cells derived from various strains of gene-targeted mice6, 7, 8.
In the second experiment, untransformed mammary epithelial cells (MCF10A) were treated with the growth inhibitory cytokine, transforming growth factor beta one (TGF-&beta;1) for 24 hours. Cells were labeled with BrdU for four hours immediately prior to harvesting. As shown in Fig. 4 BrdU labeling in S-phase cells is greatly diminished by TGF-&beta;1 signaling, the specificity of our staining is also validated with an IgG negative control (Fig. 4C). Quantification of the different phases of the cell cycle confirms that TGF-&beta;1 primarily inhibits proliferation in the G1 phase of the cell cycle, leading to an accumulation in this phase.
In our last example, pRB deficient SaOS-2 cells are transfected with a CMV-CD20 expression vector and either CMV-RB or CMV-&beta;-Gal as a control. Three days following transfection cells were harvested, stained, and fixed. Flow cytometry analysis of these cells is shown in Fig. 5. This demonstrates the cell cycle distribution of negative control transfected cells (Fig. 5A) in comparison with the distribution following 72 hours of pRB expression (Fig. 5B). Following curve fitting by Multi Cycle software, a direct comparison of the proportion of cell cycle phases is shown in Fig. 5C. This reveals the accumulation of cells in G1 following pRB expression and the relative depletion of cells from the S and G2/M phases. We have used variations on this assay to probe G1 arrest mechanisms extensively9, 10, 11, 12, 13.
These examples demonstrate how cell cycle control can be measured in response to a variety of stimuli or cell manipulations. This approach can therefore be adapted to many applications that require the measurement of cell cycle position in culture type experiments.
<img alt="Figure 1" src="/files/ftp_upload/3491/3491fig1.jpg" /> 
Figure 1. Quantitation of cell cycle phases by combined propidium iodide and BrdU staining (PI-BrdU). (A) This panel displays three dimensional flow cytometry of propidium iodide and BrdU stained cells. Note that cells with 2N and 4N DNA content are centered over the 200 and 400 marks on the X-axis scale for propidium iodide staining intensity. BrdU staining intensity is measured on a logarithmic scale on the Y-axis. Note the position of gates used to quantitate cells in the G1, S, and G2/M phases of the cell cycle. (B) The relative proportion of cells in each of the G1, S, and G2/M gates from A are shown on this graph.
<img alt="Figure 2" src="/files/ftp_upload/3491/3491fig2.jpg" /> 
Figure 2. Quantitation of cell cycle phases in selected cells using propidium iodide and the cell surface marker CD20. (A) This panel shows propidium iodide and CD20 staining for a mixture of cells, some of which ectopically express CD20 and pRB. Note that the cells with 2N and 4N DNA (the most abundant populations) are centered over the 200 and 400 marks on the X-axis. The position of the CD20 + gate selects cells that are stained at least 10X more brightly than background. (B) A graph of cell counts versus propidium iodide staining is shown for CD20 negative cells in panel A that are asynchronously proliferating. (C) A similar graph is shown for CD20 positive cells from panel A that have been induced to arrest with pRB expression and contain cells with primarily 2N DNA content. This demonstrates that an arrested sub-population can be distinguished from other cells in this culture using this staining technique.
<img alt="Figure 3" src="/files/ftp_upload/3491/3491fig3.jpg" /> 
Figure 3. Inhibition of cell proliferation by p27KIP1. (A) PI-BrdU analysis is used to measure the cell cycle phases in an asynchronously proliferating population of cells that were transduced with an empty pBABE retroviral vector. (B) A similar analysis of cells transduced with pBABE-p27. Note the absence of cells in the S-phase and greater intensity of events in G1 and G2/M gates. (C) Quantitation of cells in the respective phases of the cell cycle in asynchronously proliferating control and p27 expressing cells.
<img alt="Figure 4" src="/files/ftp_upload/3491/3491fig4.jpg" /> 
Figure 4. Inhibition of cell proliferation by TGF-&beta;1 (A) PI-BrdU analysis of asynchronously proliferating MCF10A cells. (B) Analysis of cells treated with 100 pM of TGF-&beta;1 for 24 hours. Note the accumulation of cells primarily in the G1 phase of the cell cycle. (C) Validation of BrdU staining by replacing the anti-BrdU primary antibody with a non-specific IgG control in asynchronously proliferating cells. (D) Graphical quantitation of the respective cell cycle phases from A and B.
<img alt="Figure 5" src="/files/ftp_upload/3491/3491fig5.jpg" /> 
Figure 5. Inhibition of cell proliferation by pRB. (A) Cell counts versus propidium iodide staining of CD20 and &szlig;-gal transfected cells is shown. This is an important control as transfection can partially synchronize cells, rendering the untransfected population (CD20 - cells) as an inappropriate control for the transfected sub-population. Transfected cells need to be compared with other analogously transfected cells. (B) Cell counts versus propidium iodide graph of CD20 and pRB transfected cells. Note the almost exclusive presence of a 2N peak. (C) Graphical representation of cell cycle phase proportions determined from A and B using curve fitting methods in Multi Cycle software.

<ol>
	<li>Dolbeare, F., Gratzner, H., Pallavicini, M. G., Gray, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flow+cytometric+measurement+of+total+DNA+content+and+incorporated+bromodeoxyuridine.">Flow cytometric measurement of total DNA content and incorporated bromodeoxyuridine.</a> Proc. Natl. Acad. Sci. U. S. A. 80, 5573-5573 (1983).</li><li>Givan, A. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flow+cytometry:+an+introduction.">Flow cytometry: an introduction.</a> Methods. Mol. Biol. 699, 1-1 (2011).</li><li>Kalejta, R. F., Brideau, A. D., Banfield, B. W., Beavis, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+integral+membrane+green+fluorescent+protein+marker,+Us9-GFP,+is+quantitatively+retained+in+cells+during+propidium+iodide-based+cell+cycle+analysis+by+flow+cytometry.">An integral membrane green fluorescent protein marker, Us9-GFP, is quantitatively retained in cells during propidium iodide-based cell cycle analysis by flow cytometry.</a> J. Exp. Cell. Res. 248, 322-322 (1999).</li><li>Qin, X. -. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+transcription+factor+E2F-1+is+a+downstream+target+of+RB+action.">The transcription factor E2F-1 is a downstream target of RB action.</a> Molecular and Cellular Biology. 15, 742-742 (1995).</li><li>van den Heuvel, S., Harlow, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+roles+for+cyclin-dependent+kinases+in+cell+cycle+control.">Distinct roles for cyclin-dependent kinases in cell cycle control.</a> Science. 262, 2050-2050 ( ).</li><li>Henley, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+cancer+derived+mutation+in+the+retinoblastoma+gene+with+a+distinct+defect+for+LXCXE+dependent+interactions.">A cancer derived mutation in the retinoblastoma gene with a distinct defect for LXCXE dependent interactions.</a> Cancer Cell. Int. 10, 8-8 (2010).</li><li>Francis, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+functional+connection+between+pRB+and+transforming+growth+factor+beta+in+growth+inhibition+and+mammary+gland+development.">A functional connection between pRB and transforming growth factor beta in growth inhibition and mammary gland development.</a> Mol. Cell. Biol. 29, 4455-4455 (2009).</li><li>Isaac, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+retinoblastoma+protein+regulates+pericentric+heterochromatin.">The retinoblastoma protein regulates pericentric heterochromatin.</a> Mol. Cell. Biol. 26, 3659-3659 (2006).</li><li>Julian, L. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+an+E2F1-specific+binding+domain+in+pRB+and+its+implications+for+apoptotic+regulation.">Characterization of an E2F1-specific binding domain in pRB and its implications for apoptotic regulation.</a> Oncogene. 27, 1572-1572 (2008).</li><li>Hirschi, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+overlapping+kinase+and+phosphatase+docking+site+regulates+activity+of+the+retinoblastoma+protein.">An overlapping kinase and phosphatase docking site regulates activity of the retinoblastoma protein.</a> Nat. Struct. Mol. Biol. 17, 1051-1051 (2010).</li><li>Dick, F. A., Dyson, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=pRB+contains+an+E2F1-specific+binding+domain+that+allows+E2F1-induced+apoptosis+to+be+regulated+separately+from+other+E2F+activities.">pRB contains an E2F1-specific binding domain that allows E2F1-induced apoptosis to be regulated separately from other E2F activities.</a> Mol. Cell. 12, 639-639 (2003).</li><li>Dick, F. A., Sailhamer, E., Dyson, N. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mutagenesis+of+the+pRB+pocket+reveals+that+cell+cycle+arrest+functions+are+separable+from+binding+to+viral+oncoproteins.">Mutagenesis of the pRB pocket reveals that cell cycle arrest functions are separable from binding to viral oncoproteins.</a> Mol. Cell. Biol. 20, 3715-3715 (2000).</li><li>Dick, F. A., Dyson, N. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three+regions+of+the+pRB+pocket+domain+affect+its+inactivation+by+human+papillomavirus+E7+proteins.">Three regions of the pRB pocket domain affect its inactivation by human papillomavirus E7 proteins.</a> J. Virol. 76, 6224-6224 (2002).</li></ol>

The authors have nothing to disclose.

In our experience, success with these techniques is dependent on a few key controls and experimental conditions. One is establishing an asynchronously proliferating control for use in these experiments. This control serves three important purposes. First, it ensures that culture conditions used for all experimental samples are adequate to support continual proliferation in the absence of treatment. This sample also serves the purpose of a positive control for the staining methodology to ensure that BrdU or CD20 positive cells can be detected when they are present. Lastly, this sample is used to calibrate the flow cytometer. This control sample can be used to adjust propidium iodide staining intensity to detect 2N and 4N cells at 200 and 400 respectively. Furthermore, detection sensitivity of BrdU or CD20 can be adjusted so that negative and positive signals are centered as shown in Figures 1A and 2A. When all samples have a similar concentration of cells in PI-RNase solution, then few adjustments to the cytometer are needed as subsequent samples are run. This is important for reasons shown in Fig. 4B and 5B where strong G1 accumulation creates essentially a single G1 peak that could be misinterpreted as G2/M.
The representative experiments also make other important points. First of all, they demonstrate that BrdU uptake and labeling can vary between cell types. For this reason it is important to empirically determine the length of pulse and staining conditions needed to adequately detect cells in S-phase. In general, cell types that double in culture in 24 hours or less can be labeled with BrdU in an hour. Slower growing cell types may require longer pulses and alterations to antibody staining conditions and duration. MCF10A cells are an excellent example in this regard as they were labeled with BrdU for four hours and stained with twice the standard concentration of antibodies for four times as long. Investigators should be careful not to exceed 6 hours of BrdU labeling even with very slowly growing cells as this can erroneously lead to inclusion of G2/M cells in the S-phase population. Secondly, in comparing the effects of p27KIP1 expression with those of TGF-&beta;1, it is clear that p27 can induce an accumulation in either G1 or G2/M phases while TGF-&beta;1 signaling induces a G1 arrest. Traditional means of detecting proliferation such as 3H-Thymidine incorporation and scintillation counting are unable to distinguish these possibilities.
In our alternate protocol we demonstrate the detection of a sub-population of cells in a larger untransfected population. It is important to empirically determine the most appropriate reporter for detecting transfected cells. In our experience some cell types either express cell surface markers poorly, or can&prime;t properly traffic them to the plasma membrane, resulting in an inability to detect transfected cells. Likewise, not all cells tolerate the membrane bound form of GFP. Selection of the most appropriate reporter should be made by assessing transfection efficiency of the reporter as well as the relative intensity of expression compared to untransfected controls. Ideally, heterologous expression of these molecules will be well tolerated and this is suggestive that the markers have little to no effect on cell cycle distribution.
One limitation of these types of flow cytometry approaches is that they are only able to establish relative abundances of cell cycle phases compared to one another. For this reason it is ambiguous if expression of p27 in Figure 3 truly induces an arrest in G1 and G2/M, or if it just slows progression through these phases relative to S-phase. These possibilities can be investigated further by treating a parallel sample of cells with a mitotic inhibitor such as nocodazole, or G1/S inhibitor like aphidicolin. Since these drugs create a dominant arrest in M-phase or early S-phase respectively, slowly proliferating cells will accumulate at the drug induced arrest point. For example cells arrested in G1 because of pRB expression will remain in G1 despite nocodazole treatment while control cells will accumulate in M-phase13.
Taken together, these experimental approaches offer a flexible methodology that can be applied to a wide range of mammalian cell cycle research questions. They can readily detect alterations in cell cycle progression and quantify differences when compared with controls.

<table border="1">
<tbody>
 <tr>
 <th>Name of the reagent/equipment</th>
 <th>Company</th>
 <th>Catalogue number</th>
 <th>Comments (optional)</th>
 </tr>
 <tr>
 <td>Cell Proliferation Labeling reagent</td>
 <td>GE Amersham</td>
 <td>RPN201</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Anti-BrdU Antibodies</td>
 <td>BD Biosciences</td>
 <td>347580</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Anti-Mouse FITC conjugated Antibodies</td>
 <td>Vector Labs</td>
 <td>FI-2000</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Cell Strain Filters</td>
 <td>BD Falcon</td>
 <td>352235</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Anti-CD20 Antibodies</td>
 <td>BD Biosciences</td>
 <td>347673</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Multi Cycle Software</td>
 <td>Phoenix Flow Systems</td>
 <td>N/A</td>
 <td>&nbsp;</td>
 </tr>
</tbody>
</table>

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.

Watch this Scientific Journal Video about Analysis of Cell Cycle Position in Mammalian Cells at JoVE.com

Analysis of Cell Cycle Position in Mammalian Cells

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 ...

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59.8K Views.  University of Western Ontario. 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.

Video: Analysis of Cell Cycle Position in Mammalian Cells

Long-term Imaging Mammalian Cells using Wide-Field Microscopy

Actually what I'm gonna do, mount it in here first and then swap out You. So now I'm gonna aspirate try and be as gentle as possible.Okay. A few seconds just holding.Okay.Alright.So the important features of our lifestyle imaging grid are that, so it's an inverted scope, which means the objective, the samples here in the objective is underneath. So the objective is pointing up and the whole scope, or practically the whole, practically the whole scope, including the objectives and the sample are at 37 degrees. So everything in this plastic box is at 37 degrees.And right on top of the stage we have an air and CO2 mix. That is 5%CO2. So the idea is that the sample is actually in a cell, it's like a cell incubator.And so we have, this is our mixer for the air and the CO2 and, and it gets humidified and heated and then piped onto the, the sample on the stage.Okay. And today we're just gonna be doing fluorescence one channel GFP fluorescence. This is the hose that the CO2 and 5%CO2 goes in and it just fits into the stage holder.That's important. And then we can close these, take these down, and then we go to our first position. And what I'm gonna do is go to the first field, which is my control sample, and I'll pick positions based on what I see on the screen.So I just make sure I actually look into the microscope in each new, well, because the focal plane for each of the wells is different, the, the dish is not perfectly flat on the bottom. So you will need to make fine adjustments when you move from well to the well. So right now I'm on the first, well, what I'll do is I'll focus, so I just hit, I just told, I just turned on the light, the transmitted light for the microscope.And I have make sure all the light is going to the eye pieces. And so I just look in, make sure I get a nice clean field. And then what I'll do is turn that light off, turn the fluorescence light on, make sure the cells that I'm looking at look healthy and they're not apoptotic.And, and I turn that off. And what I do next is send, so now I'm gonna start picking positions, which means I have to te send the light path to the camera. This is the camera right here.So I tell the microscope to send all the camera, they don't see anything in the eye pieces. What I'll do is in the software controlling the microscope, I'll choose the right filter set, which for our purposes is psy. And then set an exposure time.And I know these cells very well and I know that they take a 0.2 second exposure time. So I tell, that's what I tell the software to do. And what I'll do is hit focus.This is, and it's basically just alive. The, A light will turn on and you can, oh, I don't know if you can see cells, but the reason this is a good field is because there's, all the cells are, even in terms of their fluorescence, There are no apoptotic cells. Apoptotic cells tend to be very bright and there's no dirt or schmutz in the field.And you can see that they're actually, this is a metaphase cell and that's a metaphase cell, that's a cell that's probably just exited mitosis. And I can just tell that because it looks like those are, those two sets of chromatin are very nicely oriented to each other. Like they just exited an face.So that's a good field. So, and now that I know that the exposure time and the filter set is correct, and this is gonna be software specific, I'll go and I'll take positions and now the software has memorized the X, Y, Z position of that field. And then when I do pick another field, it's, and I'll, so when I'm picking, when I'm picking new fields, but in the same, well, I'll use the, I'll move the i'll, I'll turn the shutter, I'll open the shutter and I'll look to see what I see on the screen and I'll pick fields based on that versus looking in the field.So I don't see any of my tonics in there. But again, even nicely centered, these cells will go through my PS I continue, here's a good example. Here's a good example of what I would avoid that, see how bright that is?That's very bright. That's probably remnants of an apoptotic cell. And the reason I would avoid that is because since it's so bright, when the software auto focuses, it will autofocus just on that and it won't be in the same plane as all these other cells.And so I won't get any information from that field. That looks nice. There's a, this is a pro metaphase right here, That's A metaphase.So, and I'm gonna take three positions per well. So that was three positions and now I'm going to go to my next Well, so you can, so what I will probably do is take images every 10 minutes. So it's a compromise.Lifestyle imaging is always a compromise, right? You have to image, you have to keep the cells healthy, especially if you're interested in mitosis. So you have to minimize their exposure to the fluorescent line.And, but you wanna get information. So you have to take images, you have to, so you have to compromise, you have to make a choice about what you want to capture, what kind of information you wanna capture. For our purposes, mitosis tends to take about an hour.So if we're taking 10 minute intervals, that's pretty good. We see prophase, we see metaphase, we see anaphase, and we can, we can see when there are, there are treatments or drugs or siRNAs that affect those specific cell cycle stages. So for our purposes, that's enough, but we always want more.We always want more positions or finer time and interval or something like that. So it's a compromise. So, and then I'll hit start.And so our software, it does an autofocusing pass on the first pass. So once I hit start, it should start auto autofocusing and you'll see the shutter open, open and close between auto focusing. So I'll hit start.So that's the shutter opening and closing. The Z motor is engaging and it's actually trying to find the plane with the most contrast, which will be the most in focus. You can actually see on the screen going in and out of finding the best plane of focus that might be worrisome.So what it does is it, it drives the Z motor to find the best, the most in focus plane focus, well the most in image. And then it acquires picture there and then it learns that Z position. And then we'll use that Z position for the next 12 passes.And then we'll do the auto focusing pass again. What Kind of experiments do they do? Okay, so our lab is mostly interested in understanding what regulates mitic progression, or at least I'm most interested in understanding that.So with live cell imaging and especially our multi position, long-term imaging scope, you can take, you can film cells for two or three days entering anding mitosis, and you can image multiple rounds of mitosis so you can understand. And then you can put drugs or RNAs or any kind of perturbations on the cells that you would like to study. So you can understand how, how those drugs or, or genes are, are important in regulating mitotic progression.Because it's not fixed cell work and live cell work, you can understand what, when you need to have a perturbation to affect mitosis or what stage of mitosis is affected. So it's really powerful in that respect. What, what are the main technical elements of this experiments?What are the possible problems? Well, the biggest problem besides getting the rig set up, getting the rig set up is not trivial. A lot of people have the rig, but just making sure it works well for your application.The financial element is not, you know, is not small. So there's the setting up of the apparatus and then, and then once you gather the data, there's a real data analysis issue. It takes these movie, if, so, if you're gathering 40 movies each with 30 cells, so you have 40 in 40 fields that you just filmed and each of those fields has 30 cells, then you have to sit down with each one of those movies and look at each one of those cells in that movie.So it takes a really long time to do data analysis. And that's, so it takes a week to analyze data that it took you two days to collect. So there's a, that's a very big obstacle in terms of gathering lots of data.So it's not, while it's high throughput data collection, it's not high throughput analysis. So what would you make to make this experiment really high throughput? So our lab is in the process of developing a program that does the data analysis automatically.And it's a pretty decent program. It's not perfect. The human eye is always better, but it's getting there.So What are the interesting application of this experiment? Maybe other fields you can mention? Besides cell division?You mean for the, oh, for the live cell imaging? Yes.Well, you can do cell motility assays. You can, depending on the, the microscopy that's involved, you can, you know, look at how cells, you could look at tumor development if you really wanted to.But there, you, you're, if as long as you have the right imaging set up, you can, you're pretty much not limited in terms of anything. Our scope happens to be just a wide field and we use 20 by 20 x objective. So it's a pretty low resolution set up, but that's sufficient for what we need to see.So there are, but there definitely aren't, you wouldn't be limited. Anything that has any dynamic kind of experiment, which is biology, it would be, would really benefit from lifestyle imaging. So calcium imaging, although you couldn't do it on our apparatus, that's a live cell.You definitely need live cell imaging for that. Like I said, patient assays, motility assays, those kinds of things would really benefit from live cell imaging. Would you like to tell a few words about your previous experience?How long do you work on these techniques? What do you you work on before? Wow.So in my PhD I didn't work.I did a lot of confocal microscopy, but no lifestyle imaging. And so during my postdoc I was, I helped Dr.King get this apparatus that, that we just used to set up these fragment, get that up and running. And, and so during my postdoc, I had a lot of experience with both widefield with TimeLapse imaging on our scope, which is the widefield and also TimeLapse imaging with confocal microscopy.And I would say that it's really powerful. You get a lot of information, but it's not easy to get working perfectly. So it really requires a lot of attention and a lot of attention to details at every little step of the process.And, and then you still have problems. So it's a tough application, but you can really get, if, if it's what you need to have to, to study your problem, your scientific problem, then it, it's invaluable in that respect, but it's tough. So find someone who knows what they're doing and use them.That's my advice. So good.

Pulse-chase Analysis of N-linked Sugar Chains from Glycoproteins in Mammalian Cells

Attachment of the Glc3Man9GlcNAc2 precursor oligosaccharide to nascent polypeptides in the ER is a common modification for secretory proteins. Although this modification was implicated in several biological processes, additional aspects of its function are emerging, with recent evidence of its role in the production of signals for glycoprotein quality control and trafficking. Thus, phenomena related to N-linked glycans and their processing are being intensively investigated. Methods that have been recently developed for proteomic analysis have greatly improved the characterization of glycoprotein N-linked glycans. Nevertheless, they do not provide insight into the dynamics of the sugar chain processing involved. For this, labeling and pulse-chase analysis protocols are used that are usually complex and give very low yields. We describe here a simple method for the isolation and analysis of metabolically labeled N-linked oligosaccharides. The protocol is based on labeling of cells with [2-3H] mannose, denaturing lysis and enzymatic release of the oligosaccharides from either a specifically immunoprecipitated protein of interest or from the general glycoprotein pool by sequential treatments with endo H and N-glycosidase F, followed by molecular filtration (Amicon). In this method the isolated oligosaccharides serve as an input for HPLC analysis, which allows discrimination between various glycan structures according to the number of monosaccharide units comprising them, with a resolution of a single monosaccharide. Using this method we were able to study high mannose N-linked oligosaccharide profiles of total cell glycoproteins after pulse-chase in normal conditions and under proteasome inhibition. These profiles were compared to those obtained from an immunoprecipitated ER-associated degradation (ERAD) substrate. Our results suggest that most NIH 3T3 cellular glycoproteins are relatively stable and that most of their oligosaccharides are trimmed to Man9-8GlcNAc2. In contrast, unstable ERAD substrates are trimmed to Man6-5GlcNAc2 and glycoproteins bearing these species accumulate upon inhibition of proteasomal degradation.

We describe a method for analysis of the alteration of N-linked glycans through the early life of glycoproteins after their biosynthesis in mammalian cells. This is achieved by pulse-chase analysis of metabolically labeled glycans, enzymatic release from glycoproteins and examination by HPLC.

Attachment of the Glc3Man9GlcNAc2 precursor oligosaccharide to nascent polypeptides in the ER is a common modification for secretory proteins. Although ...

Analysis of DNA Double-strand Break (DSB) Repair in Mammalian Cells

DNA double-strand breaks are the most dangerous DNA lesions that may lead to massive loss of genetic information and cell death. Cells repair DSBs using two major pathways: nonhomologous end joining (NHEJ) and homologous recombination (HR). Perturbations of NHEJ and HR are often associated with premature aging and tumorigenesis, hence it is important to have a quantitative way of measuring each DSB repair pathway. Our laboratory has developed fluorescent reporter constructs that allow sensitive and quantitative measurement of NHEJ and HR. The constructs are based on an engineered GFP gene containing recognition sites for a rare-cutting I-SceI endonuclease for induction of DSBs. The starting constructs are GFP negative as the GFP gene is inactivated by an additional exon, or by mutations. Successful repair of the I-SceI-induced breaks by NHEJ or HR restores the functional GFP gene. The number of GFP positive cells counted by flow cytometry provides quantitative measure of NHEJ or HR efficiency.

Analysis of DNA Double-strand Break DSB Repair in Mammalian Cells

This article describes GFP-based fluorescence in vivo assays that separately quantify homologous recombination and nonhomologous end joining in mammalian cells.

DNA double-strand breaks are the most dangerous DNA lesions that may lead to massive loss of genetic information and cell death.  Cells repair DSBs using ...

MISSION esiRNA for RNAi Screening in Mammalian Cells

RNA interference (RNAi) is a basic cellular mechanism for the control of gene expression. RNAi is induced by short double-stranded RNAs also known as small interfering RNAs (siRNAs). The short double-stranded RNAs originate from longer double stranded precursors by the activity of Dicer, a protein of the RNase III family of endonucleases. The resulting fragments are components of the RNA-induced silencing complex (RISC), directing it to the cognate target mRNA. RISC cleaves the target mRNA thereby reducing the expression of the encoded protein1,2,3. RNAi has become a powerful and widely used experimental method for loss of gene function studies in mammalian cells utilizing small interfering RNAs.
Currently two main methods are available for the production of small interfering RNAs. One method involves chemical synthesis, whereas an alternative method employs endonucleolytic cleavage of target specific long double-stranded RNAs by RNase III in vitro. Thereby, a diverse pool of siRNA-like oligonucleotides is produced which is also known as endoribonuclease-prepared siRNA or esiRNA. A comparison of efficacy of chemically derived siRNAs and esiRNAs shows that both triggers are potent in target-gene silencing. Differences can, however, be seen when comparing specificity. Many single chemically synthesized siRNAs produce prominent off-target effects, whereas the complex mixture inherent in esiRNAs leads to a more specific knockdown10.
In this study, we present the design of genome-scale MISSION esiRNA libraries and its utilization for RNAi screening exemplified by a DNA-content screen for the identification of genes involved in cell cycle progression. We show how to optimize the transfection protocol and the assay for screening in high throughput. We also demonstrate how large data-sets can be evaluated statistically and present methods to validate primary hits. Finally, we give potential starting points for further functional characterizations of validated hits.

Here we use a human esiRNA library in a high-throughput screen for genes involved in cell division. We demonstrate how to set up and conduct an esiRNA screens, as well as how to analyze and validate the results.

RNA interference (RNAi) is a basic cellular mechanism for the control of gene expression. RNAi is induced by short double-stranded RNAs also known as ...

Mutagenesis and Functional Analysis of Ion Channels Heterologously Expressed in Mammalian Cells

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying potassium (referred to as GIRK) channels, which are important for regulating the excitability of neurons. There are four different mammalian GIRK channel subunits (GIRK1-GIRK4) - we focus on GIRK2 because it forms a homotetramer. Stimulation of different types of G protein-coupled receptors (GPCRs), such as the muscarinic receptor (M2R), leads to activation of GIRK channels. Alcohol also directly activates GIRK channels. We will show how to mutate one amino acid by specifically changing one or more nucleotides in the cDNA for the GIRK channel. This mutated cDNA sequence will be amplified in bacteria, purified, and the presence of the point mutation will be confirmed by DNA sequencing. The cDNAs for the mutated and wild-type GIRK channels will be transfected into human embryonic kidney HEK293T cells cultured in vitro. Lastly, whole-cell patch-clamp electrophysiology will be used to study the macroscopic potassium currents through the ectopically expressed wild-type or mutated GIRK channels. In this experiment, we will examine the effect of a L257W mutation in GIRK2 channels on M2R-dependent and alcohol-dependent activation.

We will demonstrate how to study the effect of a single point mutation on the function of an ion channel.

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying ...

Analysis of mRNA Nuclear Export Kinetics in Mammalian Cells by Microinjection

In eukaryotes, messenger RNA (mRNA) is transcribed in the nucleus and must be exported into the cytoplasm to access the translation machinery. Although the nuclear export of mRNA has been studied extensively in Xenopus oocytes1 and genetically tractable organisms such as yeast2 and the Drosophila derived S2 cell line3, few studies had been conducted in mammalian cells. Furthermore the kinetics of mRNA export in mammalian somatic cells could only be inferred indirectly4,5. In order to measure the nuclear export kinetics of mRNA in mammalian tissue culture cells, we have developed an assay that employs the power of microinjection coupled with fluorescent in situ hybridization (FISH). These assays have been used to demonstrate that in mammalian cells, the majority of mRNAs are exported in a splicing dependent manner6,7, or in manner that requires specific RNA sequences such as the signal sequence coding region (SSCR) 6. In this assay, cells are microinjected with either in vitro synthesized mRNA or plasmid DNA containing the gene of interest. The microinjected cells are incubated for various time points then fixed and the sub-cellular localization of RNA is assessed using FISH. In contrast to transfection, where transcription occurs several hours after the addition of nucleic acids, microinjection of DNA or mRNA allows for rapid expression and allows for the generation of precise kinetic data.

Here we describe an assay that employs the power of microinjection coupled with fluorescent in situ hybridization in order to accurately measure the nuclear export kinetics of mRNA in mammalian somatic cells.

In eukaryotes, messenger RNA (mRNA) is transcribed in the nucleus and must be exported into the cytoplasm to access the translation machinery. Although ...

Expression Analysis of Mammalian Linker-histone Subtypes

Linker histone H1 binds to the nucleosome core particle and linker DNA, facilitating folding of chromatin into higher order structure. H1 is essential for mammalian development1 and regulates specific gene expression in vivo2-4. Among the highly conserved histone proteins, the family of H1 linker histones is the most heterogeneous group. There are 11 H1 subtypes in mammals that are differentially regulated during development and in different cell types. These H1 subtypes include 5 somatic H1s (H1a-e), the replacement H10, 4 germ cell specific H1 subtypes, and H1x5. The presence of multiple H1 subtypes that differ in DNA binding affinity and chromatin compaction ability6-9 provides an additional level of modulation of chromatin function. Thus, quantitative expression analysis of individual H1 subtypes, both of mRNA and proteins, is necessary for better understanding of the regulation of higher order chromatin structure and function.
Here we describe a set of assays designed for analyzing the expression levels of individual H1 subtypes (Figure 1). mRNA expression of various H1 variant genes is measured by a set of highly sensitive and quantitative reverse transcription-PCR (qRT-PCR) assays, which are faster, more accurate and require much less samples compared with the alternative approach of Northern blot analysis. Unlike most other cellular mRNA messages, mRNAs for most histone genes, including the majority of H1 genes, lack a long polyA tail, but contain a stem-loop structure at the 3' untranslated region (UTR)10. Therefore, cDNAs are prepared from total RNA by reverse transcription using random primers instead of oligo-dT primers. Realtime PCR assays with primers specific to each H1 subtypes (Table 1) are performed to obtain highly quantitative measurement of mRNA levels of individual H1 subtypes. Expression of housekeeping genes are analyzed as controls for normalization. 
The relative abundance of proteins of each H1 subtype and core histones is obtained through reverse phase high-performance liquid chromatography (RP-HPLC) analysis of total histones extracted from mammalian cells11-13. The HPLC method and elution conditions described here give optimum separations of mouse H1 subtypes. By quantifying the HPLC profile, we calculate the relative proportion of individual H1 subtypes within H1 family, as well as determine the H1 to nucleosome ratio in the cells.

We describe a set of assays to analyze expression levels of H1 linker histones. mRNA of individual H1 genes are quantitatively measured by random primer based reverse transcription followed by real-time PCR, whereas protein quantification of H1 histones is achieved by HPLC analysis.

Linker histone H1 binds to the nucleosome core particle and linker DNA, facilitating folding of chromatin into higher order structure.  H1 is essential ...

Visualization of Endoplasmic Reticulum Localized mRNAs in Mammalian Cells

In eukaryotes, most of the messenger RNAs (mRNAs) that encode secreted and membrane proteins are localized to the surface of the endoplasmic reticulum (ER). However, the visualization of these mRNAs can be challenging. This is especially true when only a fraction of the mRNA is ER-associated and their distribution to this organelle is obstructed by non-targeted (i.e. "free") transcripts. In order to monitor ER-associated mRNAs, we have developed a method in which cells are treated with a short exposure to a digitonin extraction solution that selectively permeabilizes the plasma membrane, and thus removes the cytoplasmic contents, while simultaneously maintaining the integrity of the ER. When this method is coupled with fluorescent in situ hybridization (FISH), one can clearly visualize ER-bound mRNAs by fluorescent microscopy. Using this protocol the degree of ER-association for either bulk poly(A) transcripts or specific mRNAs can be assessed and even quantified. In the process, one can use this assay to investigate the nature of mRNA-ER interactions.

Here we describe a method to visualize endoplasmic reticulum-associated mRNAs in mammalian tissue culture cells. This technique involves the selective permeabilization of the plasma membrane with digitonin to remove cytoplasmic contents followed by fluorescent in situ hybridization to detect either bulk poly(A) mRNA or specific transcripts.

In eukaryotes, most of the messenger RNAs (mRNAs) that encode secreted and membrane proteins are localized to the surface of the endoplasmic reticulum ...

Measurement of Heme Synthesis Levels in Mammalian Cells

Heme serves as the prosthetic group for a wide variety of proteins known as hemoproteins, such as hemoglobin, myoglobin and cytochromes. It is involved in various molecular and cellular processes such as gene transcription, translation, cell differentiation and cell proliferation. The biosynthesis levels of heme vary across different tissues and cell types and is altered in diseased conditions such as anemia, neuropathy and cancer. This technique uses [4-14C] 5-aminolevulinic acid ([14C] 5-ALA), one of the early precursors in the heme biosynthesis pathway to measure the levels of heme synthesis in mammalian cells. This assay involves incubation of cells with [14C] 5-ALA followed by extraction of heme and measurement of the radioactivity incorporated into heme. This procedure is accurate and quick. This method measures the relative levels of heme biosynthesis rather than the total heme content. To demonstrate the use of this technique the levels of heme biosynthesis were measured in several mammalian cell lines.

Altered intracellular heme levels are associated with common diseases such as cancer. Thus, there is a need to measure heme biosynthesis levels in diverse cells. The goal of this protocol is to provide a fast and sensitive method to measure and compare the levels of heme synthesis in different cells.

Heme serves as the prosthetic group for a wide variety of proteins known as hemoproteins, such as hemoglobin, myoglobin and cytochromes. It is involved in ...

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, electrical excitability and cell proliferation. The diversity and specificity of Ca2+ signaling derives from mechanisms by which Ca2+ signals are generated to act over different time and spatial scales, ranging from cell-wide oscillations and waves occurring over the periods of minutes to local transient Ca2+ microdomains (Ca2+ puffs) lasting milliseconds. Recent advances in electron multiplied CCD (EMCCD) cameras now allow for imaging of local Ca2+ signals with a 128 x 128 pixel spatial resolution at rates of &#62;500 frames sec-1 (fps). This approach is highly parallel and enables the simultaneous monitoring of hundreds of channels or puff sites in a single experiment. However, the vast amounts of data generated (ca. 1 Gb per min) render visual identification and analysis of local Ca2+ events impracticable. Here we describe and demonstrate the procedures for the acquisition, detection, and analysis of local IP3-mediated Ca2+ signals in intact mammalian cells loaded with Ca2+ indicators using both wide-field epi-fluorescence (WF) and total internal reflection fluorescence (TIRF) microscopy. Furthermore, we describe an algorithm developed within the open-source software environment Python that automates the identification and analysis of these local Ca2+ signals. The algorithm localizes sites of Ca2+ release with sub-pixel resolution; allows user review of data; and outputs time sequences of fluorescence ratio signals together with amplitude and kinetic data in an Excel-compatible table.

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Here we present techniques for imaging local IP3-mediated Ca2+ events using fluorescence microscopy in intact mammalian cells loaded with Ca2+ indicators together with an algorithm that automates identification and analysis of these events.

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, ...