This project was supported by grants from the National Institutes of Health (5 P20 RR016461 and 8 P20 GM103499), the National Science Foundation (EPS&#8212;0903795). Further support provided by Furman University&#8217;s Research and Professional Growth and Furman Advantage awards.

This study was carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. The animal protocol was approved by the Furman University Institutional Animal Care and Use Committee (Permit number: A3242-01). All materials and equipment used in this protocol can be found in the Table of Materials and Equipment, while buffers used can be found in the Table of Buffers and Solutions.
1. Single Cell Suspension of Lymphocytes from Mouse Spleens
<ol>
	<li>Sacrifice mice via CO2 asphyxiation. Confirm euthanasia by cervical dislocation. 
	Note: Use spleens from two 6-8 week old isogenic mice (e.g.,&#160;C57B/6) of the same gender. Typically, two spleens are processed.</li>
	<li>Spray the animals profusely with 70% ethanol solution and orient such that the head is facing left.</li>
	<li>Using forceps, lift the skin of the animal upwards and away from the body. Using scissors, cut a small notch through the skin near the abdomen of the animal. Using fingers, pull each side of the notch to expose the peritoneum from the neck to the beginning of the hind legs. The spleen should be visible directly underneath the peritoneum.</li>
	<li>Using forceps, lift the peritoneum and make a small incision to expose the spleen. Remove the spleen with forceps and use scissors to tease away any adipose and connective tissue.</li>
	<li>Place the spleen into a 50 ml conical tube containing 10 ml PBS supplemented with 2% Fetal Bovine Serum (hereafter referred to as PBS+2%). Keep the tube on ice until ready to begin the single cell suspension. 
	Note: Typical recovery is between 60-90 million cells per spleen, and typically, 2 million cells are needed per sample. Perform all of the following steps in a tissue culture hood.</li>
	<li>Decant the spleens into a sterile 100 mm tissue culture dish.</li>
	<li>Obtain two frosted microscope slides and hold the slides so that the two rough frosted surfaces face inwards towards one another. Wet the slides in the PBS + 2% solution poured into the petri dish.</li>
	<li>Hold the spleen between the frosted surfaces of the slides, with the edges still submerged, and grind the spleen gently by moving the slides back and forth against one another (Figure 1A). The cells will fall into the Petri dish. Continue grinding until all cells have been released and the remains of the spleen appear white (Figure 1B).</li>
	<li>Draw up the cell suspension in a pipet and filter it slowly through a 70 &#956;m filter into a new sterile 50 ml conical tube. Note that small pieces of red pulp will be present on the filter (Figure 2A).
	<ol>
		<li>If the filter rises up, gently lift it up slightly to allow the fluid to drain through and place the filter back into the 50 ml conical tube and continue, repeating this process as necessary. If processing more than 5 spleens, it may be necessary to divide into two batches and use a new filter for each.</li>
	</ol>
	</li>
	<li>Using the stopper portion of a 3 ml syringe gently press the red pulp containing the remaining cells through the strainer (Figure 2B). Make sure to press both the filter bottom and sides to ensure that all red pulp has passed through the filter (Figure 2C).</li>
	<li>Add 10 mlof PBS + 2% to the tissue culture dish and use this to wash the slides, syringe stopper, and dish to recover any remaining cells. Pass this through the same 70 &#956;m cell strainer into the 50 mL conical tube. Repeat this step as necessary.</li>
	<li>Centrifuge the filtered cell suspension at 300 x g for 5 min at 4 &#176;C.</li>
	<li>Gently decant and discard the supernatant, careful not to disturb the pellet. A trace amount of PBS + 2% will be left in the tube. Cap the tube and flick the pellet until the pellet is completely resuspended.</li>
	<li>Lyse red blood cells by adding 2 mL of ACK Lysis Buffer (0.15 M NH4Cl, 1 mM KHCO3, 0.1 mM EDTA, pH to 7.2, store at 4 &#176;C) per spleen to the conical tube and rotate and invert the tube to ensure that all cells come into contact with the ACK buffer. Gently swirl the tube for 1 min at RT.</li>
	<li>Bring the volume of the cell suspension to 50 ml by adding PBS + 2% the tube to neutralize the ACK buffer. Invert the tube 10 times and centrifuge for 5 min at 300 x g at 4 &#176;C. After the centrifugation is complete, the pellet should be white (Figure 3).</li>
	<li>Gently decant and discard the supernatant, careful not to disturb the pellet. Leave a trace amount of PBS + 2% in the tube, cap the tube, and flick the pellet to resuspend it.</li>
	<li>Add 10 ml of T cell media [10% FBS , 10 mM HEPES (pH 7.0), 2 mM GlutaMAX, 1 mM sodium pyruvate, 1X non-essential amino acids, 1x penicillin/streptomycin and 50 mM &#946;-mercaptoethanol], and further resuspend the pellet by gently pipetting up and down. Then filter the suspension through a new 70 &#956;m filter into a new 50 ml conical tube.</li>
	<li>Use another 5-10 ml of T cell media to rinse the original tube and pass this through the 70 &#956;m filter into the tube containing the rest of the filtered cells. If processing more than two spleens, the volume of T cell media can be increased to maximize recovery.</li>
	<li>Keep cells on ice until ready to be stained. Cells should be counted and viability accessed.
	<ol>
		<li>To count cells, combine 3 ml of cells with 24 ml of PBS + 2% and 3 ml of 0.4% trypan blue. Load 10 ml of this into each chamber of an improved Neubauer hemocytometer. Viability, as assessed by trypan blue exclusion, is typically between 85-95%.</li>
	</ol>
	</li>
</ol>
2. Viability and Surface Staining
<ol>
	<li>Pellet cells by centrifuging for 10 min at 300 x g at 4 &#176;C. Resuspend cells in T cell media to a concentration of 1 x 107 cells/ml. Transfer 2 million cells (100 ml) into a well of a U&#8211;bottomed 96-well tissue culture plate.</li>
	<li>Centrifuge the 96-well plate for 10 min at 300 x g at 4 &#176;C.</li>
	<li>Remove the supernatant from each well by flicking liquid within the well plate into the sink (the cell pellet will remain in the well) and dabbing the plate on a clean paper towel. Wash the cells by resuspending them in 200 &#956;l PBS followed by centrifugation at 300 x g for 10 min at 4 &#176;C. Perform all washes this way unless otherwise noted. 
	Note: Do not use PBS + 2% as it will interfere with the fixable PI stain.</li>
	<li>Remove the supernatant by flicking the plate and add 100 ml of freshly prepared commercial stain (e.g., fixable red dead cell stain) to each well.
	<ol>
		<li>Make the stain by diluting the reactive dye stock solution 1:10 in DMSO followed by a 1:10 dilution in PBS. Incubate the plate at 4 &#176;C for 30 min protected from light.</li>
	</ol>
	</li>
	<li>Centrifuge the plate for 10 min at 300 x g at 4 &#176;C. At this point forward, perform all work with the plate with the tissue culture hood light off.</li>
	<li>Remove the supernatant by flicking the plate and then add 100 ml of Fc block solution. Make Fc block solution by diluting Fc block stock solution 1:100 in PBS.</li>
	<li>Dilute antibodies to be used for surface staining (e.g.,&#160;anti-CD4 or anti-CD8) in PBS and add 100 ml to each well. Incubate the 96-well plate for 30 min at 4 &#176;C protected from light. 
	Note: Surface stain antibodies should be titered for optimal performance. Typically use a 1:200 or 1:400 dilution, per the manufacturer&#8217;s protocol.</li>
</ol>
3. Intracellular Staining for Histone H3
<ol>
	<li>Following incubation of the surface stain, wash the cells twice in PBS.</li>
	<li>After the last wash, add 100 &#956;l of 4% paraformaldehyde to each well. Incubate the 96-well plate for 5 min at RT. Make 4% paraformaldehyde by diluting 16% paraformaldehyde in PBS. Make this fresh.</li>
	<li>Wash the cells twice in PBS.</li>
	<li>Make Perm/Block solution (stock Perm solution is PBS + 2% + 0.02% Triton X-100, store at 4 &#176;C) by combining 2 ml of normal rabbit serum per 100 ml stock Perm solution. Make enough for 60 ml/sample.</li>
	<li>Add 40 ml Perm/Block to each sample well and mix well by gently pipetting up and down, taking care not to make bubbles. Incubate the plate at RT for 45 min in the dark. Save remaining Perm/Block for step 3.6.</li>
	<li>Dilute fluorescently conjugated Histone H3K4me1 antibody in Perm/Block solution reserved in step 3.5 and add 10 &#956;l of this dilution to each well. Mix by gently pipetting up and down. Incubate the 96-well plate in the dark for 1 hr at 4 &#176;C. 
	Note: Use an H3K4me1 antibody conjugated using the R-Phycoerythrin conjugation kit following the manufacturer&#8217;s instructions.</li>
	<li>Wash the cells twice in PBS + 2%.</li>
	<li>Resuspend the cells in 200 &#956;l PBS+2% and transfer the samples to FACS tubes for flow cytometry analysis. The samples can be stored at 4 &#176;C in the dark. For optimal results analyze the samples within two days. Singly stained samples can be used as flow cytometry compensation controls.</li>
</ol>

<ol>
	<li>Bravo-Ferrada, B. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Study+of+surface+damage+on+cell+envelope+assesed+by+afm+and+flow+cytometry+of+lactobacillus+plantarum+exposed+to+ethanol+and+dehydration.">Study of surface damage on cell envelope assesed by afm and flow cytometry of lactobacillus plantarum exposed to ethanol and dehydration.</a> J Appl Microbiol. , (2015).</li><li>Agola, J. O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+Bead-Based+Flow+Cytometry+for+Assaying+Rab7+GTPase+Interaction+with+the+Rab-Interacting+Lysosomal+Protein+(RILP)+Effector+Protein.">Quantitative Bead-Based Flow Cytometry for Assaying Rab7 GTPase Interaction with the Rab-Interacting Lysosomal Protein (RILP) Effector Protein.</a> Methods Mol Biol. 1298, 331-354 (2015).</li><li>Toh, W. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Application+of+flow+cytometry+to+analyze+intracellular+location+and+trafficking+of+cargo+in+cell+populations.">Application of flow cytometry to analyze intracellular location and trafficking of cargo in cell populations.</a> Methods Mol Biol. 1270, 227-238 (2015).</li><li>Rawlings, J. S., Gatzka, M., Thomas, P. G., Ihle, J. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chromatin+condensation+via+the+condensin+II+complex+is+required+for+peripheral+T-cell+quiescence.">Chromatin condensation via the condensin II complex is required for peripheral T-cell quiescence.</a> EMBO J. 30, 263-276 (2011).</li><li>Lee, M. D., Bingham, K. N., Mitchell, T. Y., Meredith, J. L., Rawlings, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+mobilization+is+both+required+and+sufficient+for+initiating+chromatin+decondensation+during+activation+of+peripheral+T-cells.">Calcium mobilization is both required and sufficient for initiating chromatin decondensation during activation of peripheral T-cells.</a> Mol Immunol. 63, 540-549 (2015).</li><li>Paul, W. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Fundamental immunology. , (2013).</li><li>Feske, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+signalling+in+lymphocyte+activation+and+disease.">Calcium signalling in lymphocyte activation and disease.</a> Nat Rev Immunol. 7, 690-702 (2007).</li><li>Isakov, N., Altman, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+kinase+C(theta)+in+T+cell+activation.">Protein kinase C(theta) in T cell activation.</a> Annu Rev Immunol. 20, 761-794 (2002).</li><li>Macian, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NFAT+proteins:+key+regulators+of+T-cell+development+and+function.">NFAT proteins: key regulators of T-cell development and function.</a> Nat Rev Immunol. 5, 472-484 (2005).</li><li>Hogan, P. G., Chen, L., Nardone, J., Rao, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transcriptional+regulation+by+calcium,+calcineurin+and+NFAT.">Transcriptional regulation by calcium, calcineurin and NFAT.</a> Genes Dev. 17, 2205-2232 (2003).</li><li>Rawlings, J. S., Rosler, K. M., Harrison, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+JAK/STAT+signaling+pathway.">The JAK/STAT signaling pathway.</a> J Cell Sci. 117, 1281-1283 (2004).</li><li>Ihle, J. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=STATs:+signal+transducers+and+activators+of+transcription.">STATs: signal transducers and activators of transcription.</a> Cell. 84, 331-334 (1996).</li><li>Moriggl, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stat5+is+required+for+IL-2-induced+cell+cycle+progression+of+peripheral+T+cells.">Stat5 is required for IL-2-induced cell cycle progression of peripheral T cells.</a> Immunity. 10, 249-259 (1999).</li><li>Kouzarides, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chromatin+modifications+and+their+function.">Chromatin modifications and their function.</a> Cell. 128, 693-705 (2007).</li></ol>

The authors declare that they have no competing financial interests.

We developed a protocol that allows for the assessment of chromatin condensation in T cells. It relies on the simple observation that Histone H3 antibodies cannot access their epitopes readily in na&#239;ve cells, but upon T cell activation, these same antibodies are able to bind to their epitopes. By comparing the MFI of Histone H3 staining between treatment groups, the relative degree of condensation or decondensation can be determined. We have used this protocol to determine relative condensation status during thymocyte development and during T cell activation4. We have also used this protocol to investigate the mechanisms that control the decondensation process5.
This protocol relies on the use of a Histone H3 antibody to detect chromatin condensation status. Since there is no global change in histone modification during T cell activation4, we are able to use an H3K4me1 antibody to assess chromatin status in this protocol. We have used antibodies raised against unmodified Histone H3; however, the signal produced was much weaker overall. In our experience, antibodies raised against modified Histone H3 work better in immunofluorescence and flow cytometry assays, while antibodies against unmodified Histone H3 work better in Western blot. It must be noted that it is also possible to use antibodies raised against other histone proteins, although we have not attempted it.
The Perm/Block and Histone H3 antibody steps are the most critical steps in the protocol. The amount of Perm/Block solution used needs to be optimized each time the stock Perm solution is made. This can be done by comparing the performance of different dilutions of stock solution. A typical experiment involves comparing chromatin status in na&#239;ve T cells to those activated for 3 hr (Figure 4). One should choose the dilution that produces the greatest change in MFI over the time period analyzed. The stock solution can be diluted in PBS + 2% to decrease Perm/Block strength by first resuspending the cells in as much as 100 ml PBS + 2% after step 3.4 and then adding the Perm/Block in step 3.5. A similar pilot experiment should be used to test different dilutions of the fluorescently conjugated Histone H3 antibody each time a new batch of antibody is labelled. These steps are especially critical to successful detection of condensation differences early in T cell activation (e.g., within 3 hr of activation). Occasionally, there are cells that do not get permeabilized and thus will not stain with the Histone H3 antibody. This may happen if the cells are not resuspended well when adding the Perm/Block or if the Perm/Block is not strong enough. These cells will appear as events pressed against the axis when visualizing a histogram of Histone H3 staining. Since these events will skew the overall MFI, these events can be omitted from analysis by gating the normal distribution of Histone H3 positive cells. Additional assistance can be found in the Troubleshooting Guide (Table 1).
The inclusion of a fixable dead cell stain allows for the assessment of cell viability in the assay. This is absolutely critical when manipulating T cell activation because certain stimuli can induce cell death. In such a case, the Histone H3 antibody can bind histones in dead cells differently than live cells, leading to misinterpretation of the results.
This protocol is designed for use in 96-well plate format, permitting a high throughput analysis of chromatin status. A spleen from a 6-8 week old female mouse will typically yield between 60-90 million cells using the protocol. Since the staining protocol requires 2 million cells per sample, one can easily assay multiple treatment groups and time points in triplicate with a single spleen on a single 96-well plate. It is possible to perform the protocol with less cells per sample; however, due to the number of centrifugation steps and the inherent loss of cells at each of these steps, it is not advisable to lower the number of cells by much. We have successfully completed the protocol with 1 million cells per sample.
We used this protocol to examine chromatin status in CD4+ T helper cells and CD8+ cytotoxic T cells. This is made possible because the protocol includes standard surface staining. The protocol could easily be adapted for the examination of chromatin in other lymphocyte subpopulations by using antibodies against population-specific surface markers. This protocol could also easily be adapted to other cell types so long as antibodies recognizing relevant surface markers are available and proper fixation/permeabilization conditions are known.&#160;

Flow cytometry is a laser-based technology developed for the analysis of multiple physical and fluorescent parameters in cell populations. This technology works as suspended cells in a stream of fluid encounter a laser, exciting fluorescent markers on or within the cells. These markers then emit light that is detected and quantified by photomultiplier tubes. While flow cytometry has traditionally been used to identify populations of cells, it has proven to be a useful technology when studying an array of cell properties including cell membrane integrity, protein-protein interactions and protein trafficking1-3. We have developed a protocol that allows this technology to be used to detect the state of chromatin in T cells as they are activated in vitro4. We have also used this protocol to investigate the mechanism of activation-induced chromatin decondensation of T cells5.&#160;
T cell activation and proliferation are critical for a proper immune response. T cells, a specific subset of lymphocytes within the immune system, are required for proper immune responses and the development of immunological memory. Activation is initiated when an antigen is presented in the context of the major compatibility complex to the T cell receptor (TCR) located on the extracellular surface of quiescent T cells (reviewed in 6). This triggers a series of dynamic and highly ordered molecular events within T cells that culminate in an increase in intracellular Ca2+ concentration7 and the nuclear translocation of transcription factors required for activation (reviewed in 8-10). Once activated, T cells gain the ability to respond to Interleukin-2 (IL-2), a potent growth factor that utilizes the JAK (Janus Kinase)/STAT (Signal Transducer and Activator of Transcription) pathway to drive clonal proliferation of activated T cells11. Briefly, IL-2 stimulation results in the phosphorylation of STAT proteins, a family of latent cytosolic transcription factors. Once phosphorylated, STAT proteins dimerize, translocate to the nucleus and drive expression of genes including those involved in cell cycle progression. In T cells, IL-2 signals via STAT5, which is required for T cell proliferation12,13.
In order to achieve clonal expansion of activated T cells, those cells that have not experienced antigen-TCR engagement (na&#239;ve T cells), must have a mechanism to ignore the potent effects of IL-2. This is achieved via the regulation of chromatin status. Na&#239;ve T cells possess a condensed chromatin that prohibits STAT5-DNA engagement in response to IL-2 stimulation. Upon activation, chromatin decondenses and STAT5 can access the promoters of target genes, permitting clonal proliferation4. Interestingly, this change in chromatin status is not dependent on epigenetic modification of histone proteins (for review, see 14) as we observed no global change in histone modification during T cell activation4.&#160;
While performing these studies, we discovered that antibodies raised against histone proteins also had difficulty accessing their epitopes in na&#239;ve cells, but that upon activation could more readily bind their epitopes4. Thus, antibody binding to histones serves as a readout for chromatin condensation status. Here we present the method to use flow cytometry to detect fluorescently conjugated Histone H3 antibodies in order to assess chromatin status in T cells. Chromatin condensation in a population of cells is measured as the mean florescence intensity (MFI) of Histone H3 staining. In the context of T cell activation, the MFI of Histone H3 staining increases, signifying the decondensation of chromatin. In addition to measuring chromatin status via intracellular Histone H3 staining, this protocol also incorporates surface staining and a fixable stain for live cells, permitting analysis of subpopulations of cells.

<table border="0" cellpadding="0" cellspacing="0" style="width:584px;" width="585">
	<tbody>
		<tr height="60">
			<td height="60" style="height:60px;width:216px;">100mm tissue culture dish</td>
			<td style="width:83px;">BD Biosciences</td>
			<td style="width:119px;">353003</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:216px;">Precleaned frosted microscope slides</td>
			<td style="width:83px;">Fisher Scientific</td>
			<td>12-550-343</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:216px;">Sterile cell strainer, 70mm nylon mesh</td>
			<td style="width:83px;">Fisher Scientific</td>
			<td>22363548</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:216px;">3mL syringe, Luer-Lok tip</td>
			<td style="width:83px;">BD</td>
			<td>309585</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:216px;">Falcon 50mL polypropylene conical tube</td>
			<td style="width:83px;">Corning</td>
			<td>352098</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:216px;">LIVE/DEAD fixable red dead cell stain kit</td>
			<td style="width:83px;">Life Technologies</td>
			<td>L23102</td>
			<td style="width:167px;">Life Technologies sells versions of this kit with different colors</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:216px;">Fc Block</td>
			<td style="width:83px;">BD Biosciences</td>
			<td>553142</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:216px;">16% paraformaldehyde</td>
			<td style="width:83px;">Electron Microscopy Sciences</td>
			<td>15710</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:216px;">Triton X-100</td>
			<td style="width:83px;">Fisher Scientific</td>
			<td>BP151-100</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:216px;">Normal rabbit serum</td>
			<td style="width:83px;">Sigma-Aldrich</td>
			<td style="width:119px;">R9133</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:216px;">Anti-H3K4me1 antibody</td>
			<td style="width:83px;">Abcam</td>
			<td>ab8895</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:216px;">R-Phycoerythrin conjugation kit</td>
			<td style="width:83px;">Abcam</td>
			<td>ab102919</td>
			<td style="width:167px;">Alternatively, the LYNX rapid RPE kit from AbD Serotec can be used</td>
		</tr>
	</tbody>
</table>

the use of flow cytometry to assess the state of chromatin in t cells

During a proper immune response, quiescent T cells become activated upon antigen presentation to their antigen-specific T cell receptor. This leads to clonal proliferation of only those T cells that bear a receptor that recognizes the antigen. Chromatin decondensation is a hallmark of T cell activation and is required for T cells to acquire the ability to proliferate after antigen engagement. This change in chromatin condensation can be detected using antibodies raised against histone proteins. These antibodies cannot bind to their epitopes in na&#239;ve T cells as well as they can in activated T cells. We describe how to simultaneously stain T cell-specific surface markers, track viability with a fixable dead cell stain, and measure chromatin status via intracellular staining of Histone H3 proteins. Stained cells are analyzed by flow cytometry and chromatin condensation status is measured as the mean fluorescence intensity (MFI) of the Histone H3 stain. Chromatin decondensation during T cell activation is demonstrated as an increase in the MFI

Lymphocytes from a C57B/6 mouse were processed into a single cell suspension according to the protocol and counted using a standard hemocytometer. Cells were seeded in triplicate at 2 x 106/ml in T-cell media in 15 ml conical tubes and left untreated, or stimulated with 1 mg/mL soluble anti-CD3 antibodies (clone 4C11) for 3 hr at 37 &#176;C in a standard tissue culture incubator. Dead cells were stained and then cells were then surface stained using FITC-CD8 and APC-CD4 antibodies according to the protocol. Histone accessibility was analyzed via flow cytometry (Figure 4). In both na&#239;ve CD4+ and CD8+ T cells, the mean fluorescent intensity (MFI) is low, signifying a condensed chromatin state. As cells are activated with anti-CD3 antibodies the MFI increases significantly (p &#60;0.001, Student t-test) indicating that chromatin has decondensed.
<img alt="Figure 1" src="/files/ftp_upload/53533/53533fig1.jpg" />
Figure 1.&#160;Technique for processing spleens into a single cell suspension using frosted microscope slides. (A) The spleen is pressed against the frosted surfaces of two microscope slides. (B) The slides are moved back and forth against each other releasing lymphocytes into a 100 mm Petri dish until the remains of the spleen are white. <a href="https://www.jove.com/files/ftp_upload/53533/53533fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53533/53533fig2.jpg" />
Figure 2. Using a syringe stopper to press remaining red pulp through a 70 mm cell strainer. (A) Red pulp remaining after a spleen is processed into a single cell suspension and passed through the 70 mm cell strainer. (B) The stopper portion of a 3 ml syringe is used to gently press remaining red pulp through the cell strainer. (C) After using the syringe, there should be virtually no red pulp left in the cell strainer. <a href="https://www.jove.com/files/ftp_upload/53533/53533fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53533/53533fig3.jpg" />
Figure 3. ACK lysis of red blood cells. (A) Centrifugation of a single cell suspension generated from a single mouse spleen prior to ACK lysis. (B) After ACK lysis of red blood cells, the cell pellet should be white. <a href="https://www.jove.com/files/ftp_upload/53533/53533fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53533/53533fig4.jpg" />
Figure 4. Representative results of the protocol. (A) Gating scheme used to analyze chromatin status in CD4+ T lymphocytes. (B and C) T cells were left untreated or activated with 1 mg/ml soluble anti-CD3 antibodies for 3 hr (in triplicate). Cells were then analyzed by flow cytometry to determine chromatin condensation in CD4+ cells (B) and CD8+ cells (C). Data are the means &#177; Standard Deviation of the mean fluorescence intensity of H3K4me1 staining. *p &#60;0.001 (Student&#8217;s t-test) <a href="https://www.jove.com/files/ftp_upload/53533/53533fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Table 1" src="/files/ftp_upload/53533/53533table1.jpg" />
Table 1: Troubleshooting Guide. A quick reference to common issues and possible solutions.

Watch this Scientific Journal Video about The Use of Flow Cytometry to Assess the State of Chromatin in T Cells at JoVE.com

The Use of Flow Cytometry to Assess the State of Chromatin in T Cells

L&#39;utilisation de la cytométrie de flux pour évaluer l&#39;état de la chromatine dans les cellules T

Flow cytometry can be utilized to assess the state of chromatin within T cells. This protocol allows scientists to interpret evidence of chromatin decondensation during T cell activation demonstrated by an increase in the mean florescence intensity (MFI) of fluorescent Histone H3 antibodies

During a proper immune response, quiescent T cells become activated upon antigen presentation to their antigen-specific T cell receptor. This leads to ...

the-use-of-flow-cytometry-to-assess-the-state-of-chromatin-in-t-cells

Research

JoVE Journal

Immunology and Infection

16.8K vues.  Furman University.  La cytométrie en flux peut être utilisé pour évaluer l'état de la chromatine dans les cellules T. Ce protocole permet d'interpréter des preuves scientifiques de décondensation de la chromatine au cours de l'activation des cellules T démontrée par une augmentation de l'intensité de fluorescence moyenne (MFI) de l'histone H3 anticorps fluorescents 

Vidéo: The Use of Flow Cytometry to Assess the State of Chromatin in T Cells

Quantitative Assessment of Immune Cells in the Injured Spinal Cord Tissue by Flow Cytometry:  a Novel Use for a Cell Purification Method

Detection of immune cells in the injured central nervous system (CNS) using morphological or histological techniques has not always provided true quantitative analysis of cellular inflammation. Flow cytometry is a quick alternative method to quantify immune cells in the injured brain or spinal cord tissue. Historically, flow cytometry has been used to quantify immune cells collected from blood or dissociated spleen or thymus, and only a few studies have attempted to quantify immune cells in the injured spinal cord by flow cytometry using fresh dissociated cord tissue. However, the dissociated spinal cord tissue is concentrated with myelin debris that can be mistaken for cells and reduce cell count reliability obtained by the flow cytometer. We have advanced a cell preparation method using the OptiPrep gradient system to effectively separate lipid/myelin debris from cells, providing sensitive and reliable quantifications of cellular inflammation in the injured spinal cord by flow cytometry. As described in our recent study (Beck &amp; Nguyen et al., Brain. 2010 Feb; 133 (Pt 2): 433-47), the OptiPrep cell preparation had increased sensitivity to detect cellular inflammation in the injured spinal cord, with counts of specific cell types correlating with injury severity. Critically, novel usage of this method provided the first characterization of acute and chronic cellular inflammation after SCI to include a complete time course for polymorphonuclear leukocytes (PMNs, neutrophils), macrophages/microglia, and T-cells over a period ranging from 2 hours to 180 days post-injury (dpi), identifying a surprising novel second phase of cellular inflammation. Thorough characterization of cellular inflammation using this method may provide a better understanding of neuroinflammation in the injured CNS, and reveal an important multiphasic component of neuroinflammation that may be critical for the design and implementation of rational therapeutic treatment strategies, including both cell-based and pharmacological interventions for SCI.

Quantification of cellular inflammation in the injured/pathological CNS by flow cytometry is complicated by lipid/myelin debris that can have similar size and granulation to cells, decreasing sensitivity/accuracy. We have advanced a cell preparation method to remove myelin debris and improve cell detection by flow cytometry in the injured spinal cord.

Évaluation quantitative des cellules immunitaires dans le tissu blessés médullaires par cytométrie en flux: une nouvelle utilisation pour un procédé de purification cellulaire

Detection of immune cells in the injured central nervous system (CNS) using morphological or histological techniques has not always provided true ...

Recherche

Immunologie et infection

Human In Vitro Suppression as Screening Tool for the Recognition of an Early State of Immune Imbalance

Regulatory T cells (Tregs) are critical mediators of immune tolerance to self-antigens. In addition, they are crucial regulators of the immune response following an infection. Despite efforts to identify unique surface marker on Tregs, the only unique feature is their ability to suppress the proliferation and function of effector T cells. While it is clear that only in vitro assays can be used in assessing human Treg function, this becomes problematic when assessing the results from cross-sectional studies where healthy cells and cells isolated from subjects with autoimmune diseases (like Type 1 Diabetes-T1D) need to be compared. There is a great variability among laboratories in the number and type of responder T cells, nature and strength of stimulation, Treg:responder ratios and the number and type of antigen-presenting cells (APC) used in human in vitro suppression assays. This variability makes comparison between studies measuring Treg function difficult. The Treg field needs a standardized suppression assay that will work well with both healthy subjects and those with autoimmune diseases. We have developed an in vitro suppression assay that shows very little intra-assay variability in the stimulation of T cells isolated from healthy volunteers compared to subjects with underlying autoimmune destruction of pancreatic &beta;-cells. The main goal of this piece is to describe an in vitro human suppression assay that allows comparison between different subject groups. Additionally, this assay has the potential to delineate a small loss in nTreg function and anticipate further loss in the future, thus identifying subjects who could benefit from preventive immunomodulatory therapy1. Below, we provide thorough description of the steps involved in this procedure. We hope to contribute to the standardization of the in vitro suppression assay used to measure Treg function. In addition, we offer this assay as a tool to recognize an early state of immune imbalance and a potential functional biomarker for T1D.

Human In Vitro Suppression as Screening Tool for the Recognition of an Early State of Immune Imbalance

Tregs are potent suppressors of the immune system. There is a lack of unique surface markers to define them, hence, definitions of Tregs are primarily functional. Here we describe an optimized in vitro assay capable of identifying immune imbalance in subjects at risk to develop T1D.

Homme In Vitro</emRépression> comme outil de dépistage pour la reconnaissance d&#39;un Etat précoce de déséquilibre immunitaire

Regulatory T cells (Tregs) are critical mediators of immune tolerance to self-antigens. In addition, they are crucial regulators of the immune response ...

Fluorescence-activated Cell Sorting for Purification of Plasmacytoid Dendritic Cells from the Mouse Bone Marrow

Fluorescence-activated cell sorting (FACS) is a technique to purify specific cell populations based on phenotypes detected by flow cytometry. This method enables researchers to better understand the characteristics of a single cell population without the influence of other cells. Compared to other methods of cell enrichment, such as magnetic-activated cell sorting (MCS), FACS is more flexible and accurate for cell separation due to the ability of phenotype detection by flow cytometry. In addition, FACS is usually capable of separating multiple cell populations simultaneously, which improves the efficiency and diversity of experiments. Although FACS has some limitations, it has been broadly used to purify cells for functional studies in both in vitro and in vivo settings. Here we report a protocol using fluorescence-activated cell sorting to isolate a very rare population of immune cells, plasmacytoid dendritic cells (pDC), with high purity from the bone marrow of lupus-prone mice for in vitro functional studies of pDC.

We report a protocol using fluorescence-activated cell sorting to isolate plasmacytoid dendritic cells (pDC) with high purity from the bone marrow of lupus-prone mice for functional studies of pDC.

Fluorescence-tri cellulaire activé pour la purification de cellules dendritiques plasmacytoïdes de la moelle osseuse de souris

Fluorescence-activated cell sorting (FACS) is a technique to purify specific cell populations based on phenotypes detected by flow cytometry. This method ...

Assessing Retinal Microglial Phagocytic Function In Vivo Using a Flow Cytometry-based Assay

Microglia are the tissue resident macrophages of the central nervous system (CNS) and they perform a variety of functions that support CNS homeostasis, including phagocytosis of damaged synapses or cells, debris, and/or invading pathogens. Impaired phagocytic function has been implicated in the pathogenesis of diseases such as Alzheimer's and age-related macular degeneration, where amyloid-&#946; plaque and drusen accumulate, respectively. Despite its importance, microglial phagocytosis has been challenging to assess in vivo. Here, we describe a simple, yet robust, technique for precisely monitoring and quantifying the in vivo phagocytic potential of retinal microglia. Previous methods have relied on immunohistochemical staining and imaging techniques. Our method uses flow cytometry to measure microglial uptake of fluorescently labeled particles after intravitreal delivery to the eye in live rodents. This method replaces conventional practices that involve laborious tissue sectioning, immunostaining, and imaging, allowing for more precise quantification of microglia phagocytic function in just under six hours. This procedure can also be adapted to test how various compounds alter microglial phagocytosis in physiological settings. While this technique was developed in the eye, its use is not limited to vision research.

Assessing Retinal Microglial Phagocytic Function In Vivo Using a Flow Cytometry-based Assay

Microglial phagocytosis is critical for the maintenance of tissue homeostasis and inadequate phagocytic function has been implicated in pathology. However, assessing microglia function in vivo is technically challenging. We have developed a simple but robust technique for precisely monitoring and quantifying the phagocytic potential of microglia in a physiological setting.

Évaluation Retinal microglial phagocytaire Fonction<em&gt; In Vivo</em&gt; L&#39;utilisation d&#39;un test basé sur la cytométrie de flux

Microglia are the tissue resident macrophages of the central nervous system (CNS) and they perform a variety of functions that support CNS homeostasis, ...

Development of Stem Cell-derived Antigen-specific Regulatory T Cells Against Autoimmunity

Autoimmune diseases arise due to the loss of immunological self-tolerance. Regulatory T cells (Tregs) are important mediators of immunologic self-tolerance. Tregs represent about 5 - 10% of the mature CD4+ T cell subpopulation in mice and humans, with about 1 - 2% of those Tregs circulating in the peripheral blood. Induced pluripotent stem cells (iPSCs) can be differentiated into functional Tregs, which have a potential to be used for cell-based therapies of autoimmune diseases. Here, we present a method to develop antigen (Ag)-specific Tregs from iPSCs (i.e., iPSC-Tregs). The method is based on incorporating the transcription factor FoxP3 and an Ag-specific T cell receptor (TCR) into iPSCs and then differentiating on OP9 stromal cells expressing Notch ligands delta-like (DL) 1 and DL4. Following in vitro differentiation, the iPSC-Tregs express CD4, CD8, CD3, CD25, FoxP3, and Ag-specific TCR and are able to respond to Ag stimulation. This method has been successfully applied to cell-based therapy of autoimmune arthritis in a murine model. Adoptive transfer of these Ag-specific iPSC-Tregs into Ag-induced arthritis (AIA)-bearing mice has the ability to reduce joint inflammation and swelling and to prevent bone loss.

We present here a method to develop functional antigen (Ag)-specific regulatory T cells (Tregs) from induced pluripotent stem cells (iPSCs) for immunotherapy of autoimmune arthritis in a murine model.

Développement de cellules souches dérivées de cellules T régulatrices spécifiques de l&#39;antigène contre Autoimmunité

Autoimmune diseases arise due to the loss of immunological self-tolerance. Regulatory T cells (Tregs) are important mediators of immunologic ...

Measurement of T Cell Alloreactivity Using Imaging Flow Cytometry

The measurement of immunological reactivity to donor antigens in transplant recipients is likely to be crucial for the successful reduction or withdrawal of immunosuppression. The mixed leukocyte reaction (MLR), limiting dilution assays, and trans-vivo delayed-type hypersensitivity (DTH) assay have all been applied to this question, but these methods have limited predictive ability and/or significant practical limitations that reduce their usefulness. Imaging flow cytometry is a technique that combines the multiparametric quantitative powers of flow cytometry with the imaging capabilities of fluorescent microscopy. We recently made use of an imaging flow cytometry approach to define the proportion of recipient T cells capable of forming mature immune synapses with donor antigen-presenting cells (APCs). Using a well-characterized mouse heart transplant model, we have shown that the frequency of in vitro immune synapses among T-APC membrane contact events strongly predicted allograft outcome in rejection, tolerance, and a situation where transplant survival depends on induced regulatory T cells. The frequency of T-APC contacts increased with T cells from mice during acute rejection and decreased with T cells from mice rendered unresponsive to alloantigen. The addition of regulatory T cells to the in vitro system reduced prolonged T-APC contacts. Critically, this effect was also seen with human polyclonally expanded, naturally occurring regulatory T cells, which are known to control the rejection of human tissues in humanized mouse models. Further development of this approach may allow for a deeper characterization of the alloreactive T-cell compartment in transplant recipients. In the future, further development and evaluation of this method using human cells may form the basis for assays used to select patients for immunosuppression minimization, and it can be used to measure the impact of tolerogenic therapies in the clinic.

This paper describes a method for measuring alloreactivity in a mixed population of T cells using imaging flow cytometry.

Mesure de cellules T alloréactivité basés sur l&#39;imagerie cytométrie en flux

The measurement of immunological reactivity to donor antigens in transplant recipients is likely to be crucial for the successful reduction or withdrawal ...

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and their important role in lung development and function, as well as their lung-localized responses to infection and inflammation. To date, no unified method for identification, isolation, and handling of alveolar macrophages from humans and mice exists. Such a method is needed for studies on these important innate immune cells in various experimental settings. The method described here, which can be easily adopted by any laboratory, is a simplified approach to harvesting alveolar macrophages from bronchoalveolar lavage fluid or from lung tissue and maintaining them in vitro. Because alveolar macrophages primarily occur as adherent cells in the alveoli, the focus of this method is on dislodging them prior to harvest and identification. The lung is a highly vascularized organ, and various cell types of myeloid and lymphoid origin inhabit, interact, and are influenced by the lung microenvironment. By using the set of surface markers described here, researchers can easily and unambiguously distinguish alveolar macrophages from other leukocytes, and purify them for downstream applications. The culture method developed herein supports both human and mouse alveolar macrophages for in vitro growth, and is compatible with cellular and molecular studies.

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

This communication describes methodologies for isolation and culture of alveolar macrophages from humans and murine models for experimental purposes.

Isolement et la Culture In Vitro de Macrophages alvéolaires murines et humaines

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and ...

Multicolor Flow Cytometry-based Quantification of Mitochondria and Lysosomes in T Cells

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular components responsible for supplying cell energy; however, excess mitochondria also produce reactive oxygen species (ROS) that could cause cell death. Therefore, the number of mitochondria must constantly be adjusted to fit the needs of the cells. This dynamic regulation is achieved in part through the function of lysosomes that remove surplus/damaged organelles and macromolecules. Hence, cellular mitochondrial and lysosomal contents are key indicators to evaluate the metabolic adjustment of cells. With the development of probes for organelles, well-characterized lysosome or mitochondria-specific dyes have become available in various formats to label cellular lysosomes and mitochondria. Multicolor flow cytometry is a common tool to profile cell phenotypes, and has the capability to be integrated with other assays. Here, we present a detailed protocol of how to combine organelle-specific dyes with surface markers staining to measure the amount of lysosomes and mitochondria in different T cell populations on a flow cytometer.

This article illustrates a powerful method to quantify mitochondria or lysosomes in living cells. The combination of lysosome- or mitochondria-specific dyes with fluorescently conjugated antibodies against surface markers allows the quantification of these organelles in mixed cell populations, like primary cells harvested from tissue samples, by using multicolor flow cytometry.

Multicolor Flow Cytometry-basé Quantification des mitochondries et des Lysosomes dans les cellules T

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular ...

Isolation of Mouse Kidney-Resident CD8+ T cells for Flow Cytometry Analysis

Tissue-resident memory T cell (TRM) is a rapidly expanding field of immunology research. Isolating T cells from various non-lymphoid tissues is one of the key steps to investigate TRMs. There are slight variations in lymphocyte isolation protocols for different organs. Kidney is an essential non-lymphoid organ with numerous immune cell infiltration especially after pathogen exposure or autoimmune activation. In recent years, multiple labs including our own have started characterizing kidney resident CD8+ T cells in various physiological and pathological settings in both mouse and human. Due to the abundance of T lymphocytes, kidney represents an attractive model organ to study TRMs in non-mucosal or non-barrier tissues. Here, we will describe a protocol commonly used in TRM-focused labs to isolate CD8+ T cells from mouse kidneys following systemic viral infection. Briefly, using an acute lymphocytic choriomeningitis virus (LCMV) infection model in C57BL/6 mice, we demonstrate intravascular CD8+ T cell labeling, enzymatic digestion, and density gradient centrifugation to isolate and enrich lymphocytes from mouse kidneys to make samples ready for the subsequent flow cytometry analysis.

Isolation of Mouse Kidney-Resident CD8+ T cells for Flow Cytometry Analysis

Following viral infection, kidney harbors a relatively large number of CD8+ T cells and offers an opportunity to study non-mucosal TRM cells. Here, we describe a protocol to isolate mouse kidney lymphocytes for flow cytometry analysis.

Isolement des cellules CD8+ T résidentes du rein de souris pour l’analyse de cytométrie de flux

Tissue-resident memory T cell (TRM) is a rapidly expanding field of immunology research. Isolating T cells from various non-lymphoid tissues is one of the ...

Analysis of SAMHD1 Restriction by Flow Cytometry in Human Myeloid U937 Cells

Sterile &#945;-motif/histidine-aspartate domain-containing protein 1 (SAMHD1) inhibits replication of HIV-1 in quiescent myeloid cells. U937 cells are widely used as a convenient cell system for analyzing SAMHD1 activity due to a low level of SAMHD1 RNA expression, leading to undetectable endogenous protein expression. Based on similar assays developed in the Stoye laboratory to characterize other retroviral restriction factors, the Bishop lab developed a two-color restriction assay to analyze SAMHD1 in U937 cells. Murine Leukaemia Virus-like particles expressing SAMHD1, alongside YFP expressed from an IRES, are used to transduce U937 cells. Cells are then treated with phorbol myristate acetate to induce differentiation to a quiescent phenotype. Following differentiation, cells are infected with HIV-1 virus-like particles expressing a fluorescent reporter. After 48 h, cells are harvested and analyzed by flow cytometry. The proportion of HIV-infected cells in the SAMHD1-expressing population is compared to that in internal control cells lacking SAMHD1. This comparison reveals a restriction ratio. SAMHD1 expression leads to a five-fold reduction in HIV infection, corresponding to a restriction ratio of 0.2. Our recent substitution of RFP for the original GFP as the reporter gene for HIV infection has facilitated flow cytometry analysis.
This assay has been successfully used to characterize the effect of amino acid substitutions on SAMHD1 restriction by transducing with viruses encoding altered SAMHD1 proteins, derived from site-directed mutagenesis of the expression vector. For example, the catalytic site substitutions HD206-7AA show a restriction phenotype of 1, indicating a loss of restriction activity. Equally, the susceptibility of different tester viruses can be determined. The assay can be further adapted to incorporate the effect of differentiation status, metabolic status, and SAMHD1 modifiers to better understand the relationship between SAMHD1, cell metabolic state, and viral restriction.

Described here is an established method to determine the extent of HIV-1 restriction by the cellular inhibitory protein SAMHD1. Human myeloid lineage U937 cells are transduced with a SAMHD1 expression vector co-expressing YFP, differentiated and then challenged with HIV-RFP. The level of restriction is determined by flow cytometry analysis.

Analyse de la restriction SAMHD1 par cytométrie en flux dans les cellules myéloïdes humaines U937

Sterile &#945;-motif/histidine-aspartate domain-containing protein 1 (SAMHD1) inhibits replication of HIV-1 in quiescent myeloid cells. U937 cells are ...