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><tbody><tr><td>100 mm tissue culture dish</td><td>BD Biosciences</td><td>353003</td><td></td></tr><tr><td>Precleaned frosted microscope slides</td><td>Fisher Scientific</td><td>12-550-343</td><td></td></tr><tr><td>Sterile cell strainer, 70 mm nylon mesh</td><td>Fisher Scientific</td><td>22363548</td><td></td></tr><tr><td>3 ml syringe, Luer-Lok tip</td><td>BD</td><td>309585</td><td></td></tr><tr><td>Falcon 50 ml polypropylene conical tube</td><td>Corning</td><td>352098</td><td></td></tr><tr><td>LIVE/DEAD fixable red dead cell stain kit</td><td>Life Technologies</td><td>L23102</td><td>Life Technologies sells versions of this kit with different colors</td></tr><tr><td>Fc Block</td><td>BD Biosciences</td><td>553142</td><td></td></tr><tr><td>16% paraformaldehyde</td><td>Electron Microscopy Sciences</td><td>15710</td><td></td></tr><tr><td>Triton X-100</td><td>Fisher Scientific</td><td>BP151-100</td><td></td></tr><tr><td>Normal rabbit serum</td><td>Sigma-Aldrich</td><td>R9133</td><td></td></tr><tr><td>Anti-H3K4me1 antibody</td><td>Abcam</td><td>ab8895</td><td></td></tr><tr><td>R-Phycoerythrin conjugation kit</td><td>Abcam</td><td>ab102919</td><td>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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.

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

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 ...

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16.8K Views.  Furman University. 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

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

Examination of Thymic Positive and Negative Selection by Flow Cytometry

A healthy immune system requires that T cells respond to foreign antigens while remaining tolerant to self-antigens. Random rearrangement of the T cell receptor (TCR) &alpha; and &beta; loci generates a T cell repertoire with vast diversity in antigen specificity, both to self and foreign. Selection of the repertoire during development in the thymus is critical for generating safe and useful T cells. Defects in thymic selection contribute to the development of autoimmune and immunodeficiency disorders1-4. 
T cell progenitors enter the thymus as double negative (DN) thymocytes that do not express CD4 or CD8 co-receptors. Expression of the &alpha;&beta;TCR and both co-receptors occurs at the double positive (DP) stage. Interaction of the &alpha;&beta;TCR with self-peptide-MHC (pMHC) presented by thymic cells determines the fate of the DP thymocyte. High affinity interactions lead to negative selection and elimination of self-reactive thymocytes. Low affinity interactions result in positive selection and development of CD4 or CD8 single positive (SP) T cells capable of recognizing foreign antigens presented by self-MHC5.
Positive selection can be studied in mice with a polyclonal (wildtype) TCR repertoire by observing the generation of mature T cells. However, they are not ideal for the study of negative selection, which involves deletion of small antigen-specific populations. Many model systems have been used to study negative selection but vary in their ability to recapitulate physiological events6. For example, in vitro stimulation of thymocytes lacks the thymic environment that is intimately involved in selection, while administration of exogenous antigen can lead to non-specific deletion of thymocytes7-9. Currently, the best tools for studying in vivo negative selection are mice that express a transgenic TCR specific for endogenous self-antigen. However, many classical TCR transgenic models are characterized by premature expression of the transgenic TCR&alpha; chain at the DN stage, resulting in premature negative selection. Our lab has developed the HYcd4 model, in which the transgenic HY TCR&alpha; is conditionally expressed at the DP stage, allowing negative selection to occur during the DP to SP transition as occurs in wildtype mice10.
Here, we describe a flow cytometry-based protocol to examine thymic positive and negative selection in the HYcd4 mouse model. While negative selection in HYcd4 mice is highly physiological, these methods can also be applied to other TCR transgenic models. We will also present general strategies for analyzing positive selection in a polyclonal repertoire applicable to any genetically manipulated mice.

We present a flow cytometry-based method to examine T cell development in vivo using genetically manipulated mice on a wildtype or T cell receptor transgenic background.

A healthy immune system requires that T cells respond to foreign  antigens while remaining tolerant to self-antigens. Random rearrangement of the  T cell ...

Rapid and Robust Analysis of Cellular and Molecular Polarization Induced by Chemokine Signaling

Cells respond to chemokine stimulation by losing their round shape in a process called polarization, and by altering the subcellular localization of many proteins. Classic imaging techniques have been used to study these phenomena. However, they required the manual acquisition of many cells followed by time consuming quantification of the morphology and the co-localization of the staining of tens of cells. Here, a rapid and powerful method is described to study these phenomena on samples consisting of several thousands of cells using an imaging flow cytometry technology that combines the advantages of a microscope with those of a cytometer. Using T lymphocytes stimulated with CCL19 and staining for MHC Class I molecules and filamentous actin, a gating strategy is presented to measure simultaneously the degree of shape alterations and the extent of co-localization of markers that are affected by CCL19 signaling. Moreover, this gating strategy allowed us to observe the segregation of filamentous actin (at the front) and phosphorylated Ezrin-Radixin-Moesin (phospho-ERM) proteins (at the rear) in polarized T cells after CXCL12 stimulation. This technique was also useful to observe the blocking effect on polarization of two different elements: inhibition of actin polymerization by a pharmacological inhibitor and expression of mutants of the Par6/atypical PKC signaling pathway. Thus, evidence is shown that this technique is useful to analyze both morphological alterations and protein redistributions.

Chemokine signaling elicits marked alterations of cellular morphology and some important redistributions of intracellular proteins. Here, a rapid and detailed protocol is provided to study these events.

Cells respond to chemokine stimulation by losing their round shape in a process called polarization, and by altering the subcellular localization of many ...

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.

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

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.

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

Flow Cytometric Characterization of Murine B Cell Development

Extensive studies have characterized the development and differentiation of murine B cells in secondary lymphoid organs. Antibodies secreted by B cells have been isolated and developed into well-established therapeutics. Validation of murine B cell development, in the context of autoimmune prone mice, or in mice with modified immune systems, is a crucial component of developing or testing therapeutic agents in mice and is an appropriate use of flow cytometry. Well established B cell flow cytometric parameters can be used to evaluate B cell development in the murine peritoneum, bone marrow, and spleen, but a number of best practices must be adhered to. In addition, flow cytometric analysis of B cell compartments should also complement additional readouts of B cell development. Data generated using this technique can further our understanding of wild type, autoimmune prone mouse models as well as humanized mice that can be used to generate antibody or antibody-like molecules as therapeutics.

We describe herein a simple analysis of the heterogeneity of the murine immune B cell compartment in the peritoneum, spleen, and bone marrow tissues by flow cytometry. The protocol can be adapted and extended to other mouse tissues.

Extensive studies have characterized the development and differentiation of murine B cells in secondary lymphoid organs. Antibodies secreted by B cells ...

A DNA/Ki67-Based Flow Cytometry Assay for Cell Cycle Analysis of Antigen-Specific CD8 T Cells in Vaccinated Mice

The cell cycle of antigen-specific T cells in vivo has been examined by using a few methods, all of which possess some limitations. Bromodeoxyuridine (BrdU) marks cells that are in or recently completed S-phase, and carboxyfluorescein succinimidyl ester (CFSE) detects daughter cells after division. However, these dyes do not allow identification of the cell cycle phase at the time of analysis. An alternative approach is to exploit Ki67, a marker that is highly expressed by cells in all phases of the cell cycle except the quiescent phase G0. Unfortunately, Ki67 does not allow further differentiation as it does not separate cells in S-phase that are committed to mitosis from those in G1 that can remain in this phase, proceed into cycling, or move into G0.
Here, we describe a flow cytometric method for capturing a &#34;snapshot&#34; of T cells in different cell cycle phases in mouse secondary lymphoid organs. The method combines Ki67 and DNA staining with major histocompatibility complex (MHC)-peptide-multimer staining and an innovative gating strategy, allowing us to successfully differentiate between antigen-specific CD8 T cells in G0, in G1 and in S-G2/M phases of the cell cycle in the spleen&#160;and draining lymph nodes of mice after vaccination with viral vectors carrying the model antigen gag of human immunodeficiency virus (HIV)-1.
Critical steps of the method were the choice of the DNA dye and the gating strategy to increase the assay sensitivity and to include highly activated/proliferating antigen-specific T cells that would have been missed by current criteria of analysis. The DNA dye, Hoechst 33342, enabled us to obtain a high-quality discrimination&#160;of the G0/G1 and G2/M DNA peaks, while preserving membrane and intracellular staining. The method has great potential to increase knowledge about T cell response in vivo and to improve immuno-monitoring analysis.

Clonal expansion is a key feature of antigen-specific T cell response. However, the cell cycle of antigen-responding T cells has been poorly investigated, partly because of technical limitations. We describe a flow cytometric method to analyze clonally expanding antigen-specific CD8 T cells in spleen and lymph nodes of vaccinated mice.

The cell cycle of antigen-specific T cells in vivo has been examined by using a few methods, all of which possess some limitations. Bromodeoxyuridine ...

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

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

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

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

Biology

Mouse Brain Histone Modification Quantification Using Flow Cytometry

Quantification of Global Histone Post Translational Modifications Using Intranuclear Flow Cytometry in Isolated Mouse Brain Microglia

Gene expression control occurs partially by modifications in chromatin structure, including the addition and removal of posttranslational modifications to histone tails. Histone post-translational modifications (HPTMs) can either facilitate gene expression or repression. For example, acetylation of histone tail lysine residues neutralizes the positive charge and reduces interactions between the tail and negatively charged DNA. The decrease in histone tail-DNA interactions results in increased accessibility of the underlying DNA, allowing for increased transcription factor access. The acetylation mark also serves as a recognition site for bromodomain-containing transcriptional activators, together resulting in enhanced gene expression. Histone marks can be dynamically regulated during cell differentiation and in response to different cellular environments and stimuli. While next-generation sequencing approaches have begun to characterize genomic locations for individual histone modifications, only one modification can be examined concurrently. Given that there are hundreds of different HPTMs, we have developed a high throughput, quantitative measure of global HPTMs that can be used to screen histone modifications prior to conducting more extensive genome sequencing approaches. This protocol describes a flow cytometry-based method to detect global HPTMs and can be conducted using cells in culture or isolated cells from in vivo tissues. We present example data from isolated mouse brain microglia to demonstrate the sensitivity of the assay to detect global shifts in HPTMs in response to a bacteria-derived immune stimulus (lipopolysaccharide). This protocol allows for the rapid and quantitative assessment of HPTMs and can be applied to any transcriptional or epigenetic regulator that can be detected by an antibody.

Author Spotlight: Enhancements in Gene Expression Regulation Research

This work describes a protocol for the quantification of global histone modifications using intranuclear flow cytometry in isolated brain microglia. The work also contains the microglia isolation protocol that was used for data collection.

Gene expression control occurs partially by modifications in chromatin structure, including the addition and removal of posttranslational modifications to ...

Neuroscience

Isolation of CD4+ T-cells and Analysis of Circulating T-follicular Helper (cTfh) Cell Subsets from Peripheral Blood Using 6-color Flow Cytometry

Aberrant T-follicular helper (Tfh) cell activity is detectable in autoimmune conditions and their presence is associated with clinical outcomes when the lymph node microenvironment in B-cell non-Hodgkin&#39;s lymphoma is analyzed. Subsets of circulating T-follicular helper cells (cTfh), the circulating memory compartment of Tfh cells in the blood, are also perturbed in disease and therefore represent potential novel predictive biomarkers. Peripheral blood-based testing is advantageous because it is relatively non-invasive and allows simple serial monitoring.This article describes a method for isolating CD4+ T-cells from human blood, and further analysis by flow-cytometry to enumerate cTfh cells and the proportions of their various subsets (cTfhPD-1-/+/hi, cTfh1,2,17 and cTfh1/17). The level of these subsets was then compared between normal subjects and patients with lymphoma. We found that the method was robust enough to obtain reliable results from routinely collected patient material. The technique we describe for the analysis can be easily adapted to cell sorting and downstream applications such as RT-PCR.

Isolation of CD4+ T-cells and Analysis of Circulating T-follicular Helper cTfh Cell Subsets from Peripheral Blood Using 6-color Flow Cytometry

Here, a protocol for the isolation and characterization of CD4+ T-cell subsets from human peripheral blood is described. Purified CD4+ T-cells are analyzed by flow cytometry to determine proportions of T-follicular helper cell subsets.

Aberrant T-follicular helper (Tfh) cell activity is detectable in autoimmune conditions and their presence is associated with clinical outcomes when the ...

A Flow Cytometry Assay to Identify Immune Cell Subsets in Peripheral Blood Mononuclear Cells

Source: Torchia, M. L. G., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/58955">Discrimination of Seven Immune Cell Subsets by Two-fluorochrome Flow Cytometry.</a> J. Vis. Exp. (2019)
This video demonstrates a flow cytometry-based technique using two fluorochromes to identify CD4+ and CD8+ T cells, &#947;&#948; T cells, B cells, natural killer (NK) cells, and monocytes in human peripheral blood mononuclear cells (PBMCs). Upon incubating the cells with fluorochrome-conjugated antibodies against the surface markers, the cell subsets are identified using an optimal flow cytometric gating strategy.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
NOTE: This protocol has been tested ...