This study was supported by the National Institute of Arthritis and Musculoskeletal and Skin Diseases, &#60;https://www.niams.nih.gov/&#62;, award number P30-AR053503; The Stabler Foundation,&#160;www.stablerfoundation.org; National Institute of Allergy and Infectious Disease, www.niaid.nih.gov, T32AI007247; Nina Ireland Program for Lung Health (NIPLH), https://pulmonary.ucsf.edu/ireland/.

All studies of human materials were approved by the Johns Hopkins Institutional Review Board under the Health Insurance Portability and Accountability Act. Patient and control samples were de-identified. PBMCs and blood from healthy controls were obtained by informed consent.
NOTE: This protocol has been tested on freshly or frozen isolated peripheral blood cells and whole blood.
1. Cell Preparation
<ol>
	<li>Isolation of peripheral blood cells (PBMC) from whole blood
	<ol>
		<li>Draw blood in a 10 mL green-top tube containing sodium heparin. After the tube has been filled with blood, immediately invert the tube several times to prevent coagulation. 
		NOTE: Tubes containing other anticoagulant such as ethylenediaminetetraacetic acid (EDTA) or sodium citrate can be used with comparable results. If a different amount of blood is collected, the following steps in the protocol should be scaled accordingly.</li>
		<li>Carefully transfer drawn blood into a 50 mL conical tube. Dilute the blood with an equal amount of phosphate buffered saline (PBS) without calcium and magnesium.</li>
		<li>Add 15 mL of density gradient medium (e.g., Ficoll) to the bottom of a new 50 mL conical tube and carefully overlay the diluted blood on top of density gradient medium, avoiding any mixing between the density gradient medium and diluted blood. 
		NOTE: This is a critical step for a proper separation of PBMCs from red cells.</li>
		<li>Centrifuge at 400 x g for 30 min at room temperature (RT), with no brake to avoid disruption of the interface.</li>
		<li>After centrifugation, carefully aspirate the upper layer using a pipette and discard it, paying attention to not remove cells at the interface between the plasma and density gradient medium, that is where PBMCs stratifies. Collect as many cells as possible from the interface without touching the red cell pellet at the bottom of the 50 mL conical tube and transfer to a new 50 mL conical tube.</li>
		<li>Add PBS to bring the final volume to 25 mL and invert several times to mix. Centrifuge at 300 x g for 10 min at RT with the brake on to remove any density gradient medium contamination from the cell suspension.</li>
		<li>Carefully aspirate supernatant without disturbing cell pellet. Add PBS to bring the final volume to 25 mL and invert several times to mix. Centrifuge at 200 x g for 10 min at RT with the brake on to remove platelets.</li>
		<li>Carefully aspirate the supernatant without disturbing the cell pellet. Resuspend PBMCs in 1 mL of PBS/0.1% sodium azide. Sodium azide reduces capping, shedding, internalization of the antibodies, and increase cell recovery, but its toxicity can impair cell viability. For this reason, sodium azide should be avoided in all steps if the cells will be cultured for subsequent experiments. Cell isolation and staining without sodium azide gave similar results to staining performed in the presence of sodium azide. 
		&#8203;CAUTION: Sodium azide can cause death by affecting the central nervous system. Contact may cause burns to skin and eyes.</li>
		<li>Dilute the cells for counting by transferring 30 &#181;L of PBMCs in a 1.5 mL microcentrifuge tube. Then add 120 &#181;L of PBS and 150 &#181;L of Trypan blue to determine cell number and viability (1:10 cell dilution). Resuspend carefully.</li>
		<li>Transfer 10 &#181;L to a hemocytometer, count the cells accordingly to the used counting chamber and determine the number of viable cells. Viable cells = number of Trypan blue negative cells total x 10 (the dilution factor used in this protocol) x 104. 
		NOTE: On average, 7-15 x 106 of PBMCs should be collected from 10 mL of blood.</li>
		<li>Resuspend the cells at 10 x 106 per mL of PBS/0.1% sodium azide 
		NOTE: PBMC can be frozen and stored in liquid nitrogen for an extended period of time. However, expression of markers such as chemokine receptors can be altered by this procedure.</li>
	</ol>
	</li>
	<li>Preparation of cells from frozen PBMC 
	NOTE: Different freezing procedures can affect cell recovery and viability12. PBMCs for these experiments were frozen in specially formulated freezing media or fetal bovine serum (FBS)/10% dimethyl sulfoxide (DMSO) with similar results. 
	&#8203;CAUTION: DMSO can be slightly hazardous in case of inhalation (lung irritant), skin contact (irritant, permeator), of eye contact (irritant), of ingestion.
	<ol>
		<li>Remove frozen PBMC from liquid nitrogen and place on ice. Transfer the cryovial from the ice directly into the 37 &#176;C water bath. Keep the cryovial at water surface and gently shake the vials until a small ice pellet remain. Transfer the cryovial back on ice.</li>
		<li>Remove any water with a 70% ethanol sprayed wipe. Slowly add 1 mL of PBS/0.1% sodium azide at 4 &#176;C and carefully transfer the cell to a 15 mL conical tube. Slowly add cold PBS/0.1% sodium azide at 4 &#176;C to reach a final volume of 15 mL.</li>
		<li>Centrifuge at 300 x g for 10 min. Carefully remove the supernatant, resuspend the cells in 1 mL of PBS/0.1% sodium azide. 
		NOTE: The cells can also be resuspended in RPMI 10% FBS with similar results.</li>
		<li>Dilute the cell for counting by transferring 30 &#181;L of PBMCs in a 1.5 mL microcentrifuge tube. Then add 120 &#181;L of PBS and 150 &#181;L of Trypan blue to determine cell number and viability (1:10 cell dilution). Resuspend carefully.</li>
		<li>Transfer 10 &#181;L to a hemocytometer, count the cells accordingly to the used counting chamber and determine the number of viable cells. Viable cells = number of Trypan blue negative cells total x 10 (the dilution factor used in this protocol) x 104.</li>
		<li>Resuspend the cells at 10 x 106 per mL in PBS/0.1% sodium azide.</li>
	</ol>
	</li>
	<li>Preparation of cells from whole blood
	<ol>
		<li>Draw blood in a 10 mL green-top tube containing sodium heparin. After the tube has been filled with blood, immediately invert the tube several times to prevent coagulation. 
		NOTE: Tubes containing other anticoagulant such as EDTA or sodium citrate can be used with comparable results. If a different amount of blood is collected, the following steps in the protocol should be scaled accordingly.</li>
		<li>Transfer 200 &#181;L of whole blood to a 12 mm x 75 mm capped tube, add 2 &#181;L of sodium azide and vortex gently for 2 s.</li>
	</ol>
	</li>
</ol>
2. Cell Staining
NOTE: Choosing pairs of fluorochromes with virtually no spectral overlap is important to reduce spread of data due to high spillover of a fluorochrome in the other fluorochrome detector. To achieve an optimal identification of al the cell subsets, fluorochromes with a high quantum yield should be used such as antibody pairs PE-BV421 and PE-APC.
<ol>
	<li>Staining of fresh and frozen PBMC
	<ol>
		<li>Transfer 100 &#181;L of PBMC (1 x 106 cells) to a 96-well V-bottom plate. 
		NOTE: Any number of cells lower than 1 x 106 can be used with similar results2.</li>
		<li>Centrifuge at 350 x g for 3 min at RT and carefully aspirate the supernatant without disturbing the cell pellet. Add to each well 100 &#181;L of PBS containing a live/dead fixable dye that reacts with free amine on proteins for 10 min to label dead cells.</li>
		<li>Prepare for each sample 30 &#181;L of a mix containing all the antibodies (anti-CD3. -CD56, TCR&#947;&#948; in fluorchrome A, and anti-CD4, CD8, CD19, CD14 in fluorchrome B). Concentration of antibodies are indicated in Table 1 and Table2. At this stage, titrated antibodies against different target molecules and in different fluorochromes can be added as well (e.g., Table 3). 
		NOTE: Antibodies concentration can vary depending on the manufacturer and lot number. Therefore, preliminary tests should be done to achieve the optimal signal. PBS, PBS/0.5% BSA, PBS/0.2% sodium azide or PBS/0.5% BSA/0.1% sodium azide were used with similar results to dilute antibodies2. Cell can also be stained in volumes different from 30 &#181;L to easily integrate this methodology to already existing staining protocols.</li>
		<li>Centrifuge at 350 x g for 3 min at RT and carefully aspirate the supernatant without disturbing the cell pellet. Add the antibody cocktail to each well and resuspend carefully without generating bubbles. Incubate for 30 min at RT in the dark. 
		NOTE: It is possible to stain the samples at 4 &#176;C with similar results.</li>
		<li>Add 150 &#181;L of staining buffer and centrifuge at 350 x g for 3 min at RT and carefully aspirate the supernatant without disturbing the cell pellet. Resuspend the cells in 200 &#181;L of PBS and acquire data on a flow cytometer. If staining volumes are changed, please ensure at least a 20-fold dilution of the original antibody mix used to wash the excess of antibodies. 
		NOTE: Stained cells can be fixed with in PBS/2% paraformaldehyde, kept in a refrigerator at 4 &#176;C overnight and then acquired on a flow cytometer the following day. 
		CAUTION: Paraformaldehyde is harmful if swallowed and can cause skin irritation</li>
	</ol>
	</li>
	<li>Staining of whole blood
	<ol>
		<li>To each 12 mm x 75 mm capped tube, add the cocktail of antibodies (anti-CD3. -CD56, TCR&#947;&#948; in fluorochrome A, and anti-CD4, CD8, CD19, CD14 in fluorochrome B) and incubate at RT for 30 min at RT in the dark. Concentration of antibodies are indicated in Table 4. At this stage, titrated antibodies against different target molecules and in different fluorochromes can be added as well. Antibody concentration can vary depending on the manufacturer and lot number. Therefore, preliminary tests should be done to achieve the optimal signal. 
		NOTE: It is possible to stain samples at 4 &#176;C with similar results.</li>
		<li>Centrifuge at 350 x g for 3 min at RT and carefully aspirate the supernatant without disturbing the cell pellet. Make a 1x solution of the red blood cell lysis buffer following manufacturer&#8217;s instructions. 
		NOTE: The lysis solution should be at RT before use.</li>
		<li>Add 2.0 mL of 1x red blood cell lysis buffer to each tube, cap well and invert several times to mix. Cover with foil and let sit for 15 min.</li>
		<li>Spin down at 300 x g for 5 min. Carefully aspirate the supernatant without disturbing the cell pellet.</li>
		<li>Add 2 mL of PBS and centrifuge at 300 x g for 5 min. 
		NOTE: At this point, the cell pellet should have a pale white coloration indicating a successful red blood cell lysis. Carefully aspirate the supernatant to not disturb the cell pellet and gently resuspend the cells in 200 &#181;L of PBS.</li>
		<li>Strain the cells through 12 mm x 75 mm tubes with 40 &#181;m filter caps to remove cell aggregates and acquire data on a flow cytometer.</li>
	</ol>
	</li>
</ol>
3. Antibody Titration
NOTE: Antibody titration is the most critical step for obtaining high-quality, reproducible data. Titration of anti-CD3, -CD8, -CD14, -CD19 and -TCR &#947;&#948; follows the standard procedure by which the concentration of antibody to optimally separate positive and negative peaks is derived by maximum staining index13,14. Dilutions at the peak or closer to the peak on the rising side of the stain index curve should be selected (Figure 1A-C). The anti-CD4 antibody is titrated to place the peak of the CD4 positive population between CD3 single positive populations and CD3+/CD8+ T cells, closer to the CD3 single positive signal to better discriminate the CD8dim populations (CD8+ &#947;&#948; T cells and NK T cells). Along the same line, CD56 titration aims to position NK CD56+ cells between the CD3+ and the CD3- population.
<ol>
	<li>Maximum stain index curve 
	NOTE: Titration of anti-CD3, -CD8, -CD14, -CD19 and -TCR &#947;&#948; follows the standard procedure by which the concentration of antibody to optimally separate positive and negative peaks is derived by a maximum staining index curve15. If antibodies against other markers are added to the panel, they also need to be titrated with a maximum staining index curve.
	<ol>
		<li>Prepare a 2-fold antibody dilution by filling 10 wells of a 96-well plate with 40 &#181;L of staining buffer. In the first well, increase the final volume to 80 &#181;L of staining buffer and add the antibody of interest at a concentration 4 times the concentration suggested by the manufacturer.</li>
		<li>Mix well and transfer 40 &#181;L to the second well. Mix well and repeat this step for the all the other wells.</li>
		<li>Stain 10 samples of PBMC or whole blood with 30 &#181;L of the 10 different 2-fold dilutions of antibodies following the protocol described before.</li>
		<li>Acquire data with a flow cytometer and plot the signal from each dilution (Figure 1A).</li>
		<li>Gate on the negative and positive populations for each antibody concentration. Increasing concentration of antibodies can lead to a higher background. Therefore, resize the negative gate accordingly.</li>
		<li>For each antibody concentration, extract information about the median and standard deviation for the fluorescent intensity of the negative population, and the median for the fluorescent intensity of the positive population. Calculate for each antibody concentration the stain index with this formula: (median fluorescent intensity of the positive population &#8212; median fluorescent intensity of the negative population) &#247; (2 x standard deviation of the fluorescent intensity of the negative population) (Figure 1B).</li>
		<li>Plot the stain index vs. the antibody concentration expressed as fraction of the antibody dilution (e.g., 1:10 dilution = 0.1), and identify the concentration of antibody with the maximum stain index value (Figure 1C).</li>
	</ol>
	</li>
	<li>Anti-CD4 and -CD56 antibody titration 
	NOTE: Anti-CD4 and -CD56 antibodies titration relies on previous titration the other markers in the two-fluorochrome panel. For the anti-CD4 antibody the titration aims at placing the anti-CD4 signal between the double CD8+/CD3+ signal and the CD3 single positive population (Figure 1D).
	<ol>
		<li>Titrate the anti-CD4 and CD56 antibodies with a 2-fold dilution strategy as described before, adding additional concentrations in between to finely identify the range of concentration that allow to separate CD4+&#160;T cells and NK cells from the other cell populations.</li>
		<li>Titrate the anti-CD4 antibody by placing the anti-CD4 signal between the double CD8+/CD3+ signal and the CD3 single positive population (Figure 1D). 
		NOTE: Special care should be done to clearly separate CD4+ T cells from CD8+ dim populations.</li>
		<li>Titrate the anti-CD56 antibody following a strategy similar to the anti-CD4 antibody titration, by placing NK cells between the CD3-negative and the CD3-positive populations.</li>
	</ol>
	</li>
</ol>
4. Gating Strategy
<ol>
	<li>Identify lymphocytic and monocytic cell populations and remove dead cells and most of the residual red blood cells from the analysis.
	<ol>
		<li>Select the entire population containing lymphocytes and monocytes based on forward vs side scatter area (FSC-A vs SSC-A). Remove cell aggregates from the analysis via forward scatter height vs forward scatter width (FSC-H vs FSC-W) and side scatter height vs side scatter width (SSC-H vs SSC-W).</li>
		<li>Use a live/dead discrimination marker to exclude bright positive dead cells and residual red blood cells from the analysis. Gate on lymphocytes and monocytes based on the different FSC-A and SSC-A profile.</li>
	</ol>
	</li>
	<li>Two-fluorochrome seven-marker gating strategy of the lymphocytic populations. 
	NOTE: Within the CD3 positive subgroup, CD4+, CD8+ and &#947;&#948; T cells can be separated using antibodies that solely target CD4, CD8 and the &#947;&#948; receptor. In a comparable way, within the CD3 negative subgroup, B cells, NK cells and monocytes can be uniquely identified using antibodies against CD19, CD56 and CD14, respectively.
	<ol>
		<li>Select the lymphocyte gate and create a dot plot with on each axis one of two fluorochromes used in this protocol (Figure 2B).</li>
		<li>Gate on CD8+ T cells identified as CD3+/CD8+ double positive cells at the top right corner of the dot-plot (Figure 2B). Exclude the dim CD8 population which might contain NKT cells. Gate on CD4+ T cells identified as population in between CD8+ T cells and the CD3 single positive populations. Gate on &#947;&#948; T cells identified as high CD3 cells. Subdivide &#947;&#948; T cells in CD8 positive and CD8 negative.</li>
		<li>Gate on NK cells identified as the population in between CD3-positive and CD3-negative cells. Subdivide NK cells in CD8 positive and CD8 negative. Gate on B cells identified as CD3-negative CD19+ population on the right lower corner of the dot plot.</li>
		<li>Select the monocyte gates and create a dot plot with on each axis one of two fluorochromes used in this protocol (Figure 2C). Gate on the CD3-/CD14+ population.</li>
	</ol>
	</li>
</ol>

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+and+Phenotypic+Characterization+of+CD57+CD4++T+Cells+and+Their+Association+with+HIV-1-Induced+T+Cell+Dysfunction.">Functional and Phenotypic Characterization of CD57+CD4+ T Cells and Their Association with HIV-1-Induced T Cell Dysfunction.</a> The Journal of Immunology. 175 (12), 8415-8423 (2005).</li><li>Focosi, D., Bestagno, M., Burrone, O., Petrini, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CD57++T+lymphocytes+and+functional+immune+deficiency.">CD57+ T lymphocytes and functional immune deficiency.</a> Journal of Leukocyte Biology. 87 (1), 107-116 (2010).</li></ol>

The authors have nothing to disclose.

The protocol presented here has been shown to be quite flexible and insensitive to changes in staining buffer, temperature and peripheral blood cell preparation due to the high expression of lineage markers on the cell surface. The most critical step for obtaining high-quality, reproducible data is antibody titration. Of note, since the titration of antibodies should always be performed during the setup of a flow cytometric panel, this step does not add extra bench-time to our two-fluorochrome approach. Titration of anti...

Flow cytometry is a technique that was developed to analyze multiple parameters on single particles at a rate of several thousand of events per second1. Examples of specimens analyzed by flow cytometry include, but are not limited to, cells, beads, bacteria, vesicles and chromosomes. A fluidic system directs particles at the interrogation point where each particle intersects its path with one or more lasers, and multiple parameters are recorded for further analysis. Forward and side scatters, generated by scattering of the pure laser light, are used to identify the target population and retrieve information about the relative size and internal complexity/granularity of particles, respectively. All the other parameters, that account for most of data in a flow cytometric analysis, are derived by fluorochrome-labeled probes that recognize and bind to specific targets on the particles of interest.
Flow cytometry is a primary tool for immunological studies to identify and characterize cell populations. To dissect the complexity of the immune system, multicolor panels are constantly evolving to expand the number of markers simultaneously recorded for deep immunophenotyping of cell populations1. This is leading to the development of more capable instruments and fluorochromes, with recent high-end flow cytometers exceeding 20 fluorescent parameters. This results in complex study design due to fluorochrome spectral overlap and in higher costs associated with custom antibody labelling and skilled operators. In several instances, complexity and costs are reduced by using separate panels of markers for different cell populations. This approach, however, is error prone, reduces the information in each panel, and can be difficult to apply to samples with limited number of cells. Moreover, increasing the number of markers precludes deep immunophenotyping on instruments with fewer fluorescent parameters. We previously developed a staining protocol to identify major human immune populations (CD4+ and CD8+ T cells, &#947;&#948; T cells, B cells, NK cells and monocytes) in peripheral blood mononuclear cells (PBMCs) by combining seven lineage markers using only two fluorochromes instead of the seven required using the traditional &#34;one fluorochrome-one marker&#34; approach (www.hcdm.org)2,3.&#160;Our initial report explored and validated the notion of combining seven markers in two fluorochromes for deep immunophenotyping. In this report, we present a step-by step protocol to isolate and stain peripheral blood cells, focusing on the practical aspect and troubleshooting steps to achieve a successful staining.&#160;
This protocol is based on the observation that lineage markers have a constant expression on the cell surface and that each cell population has an exclusive combination of lineage markers. In PBMCs, CD3 expression subdivides immune cells into two main categories: CD3-positive T lymphocytes and CD3-negative cells. Within the CD3 positive subgroup, CD4+, CD8+ and &#947;&#948; T cells can be separated using antibodies that solely target CD4, CD8 and the &#947;&#948; receptor. In a comparable way, within the CD3 negative subgroup, B cells, NK cells and monocytes can be uniquely identified using antibodies against CD19, CD56 and CD14, respectively. In a standard one fluorochrome-one marker approach, anti-CD3, -CD4, -CD8, -CD14, -CD19, -CD56 and -TCR &#947;&#948; antibodies are detected with seven different fluorochromes. Our approach combines anti-CD3, -CD56, and -TCR &#947;&#948; antibodies in one fluorochrome (labeled for convenience fluorochrome A) and anti-CD4, -CD8, -CD14 and -CD19 antibodies in a different fluorochrome (fluorochrome B). This is possible by a combination of antibody titration and differential antigen expression. Both CD4+ and CD8+ T cells are positive for the anti-CD3 antibody in fluorochrome A, but they can be separated in fluorochrome B maximizing the expression of the CD8 signal while placing, with an ad hoc titration, the CD4 signal in between the CD8 and the CD3 positive-CD4/CD8 double negative cells. &#947;&#948; T cells expresses higher level of CD3 than CD4 and CD8, and therefore they can be identified as CD3 high4. This signal is further boosted by labeling &#947;&#948; T cells in fluorochrome A with an anti-TCR &#947;&#948; antibodies, thus improving separation between CD3 low T cells and CD3 high &#947;&#948; T cells. B cells can be identified as CD3- in fluorochrome A and CD19+ in fluorochrome B. To separate CD3 negative NK cells from B cells, an anti-CD56 antibody was used in fluorochrome A as the anti-CD3. This is possible because CD56 expressed on NK cell at a much lower level than CD3 on T cells5. Finally, monocytes can be identified via a combination of forward-side scatter properties and expression of CD14 in fluorochrome B.
The idea of combining up to four markers using two fluorochromes has been already successfully attempted before6,7,8, and has been used in a clinical protocol to identify malignant lymphocytic populations9. A previous report also combined seven markers (with different specificity from the markers than we used in our protocol) using two fluorochromes, but this approach relied on a complex labelling of each antibody with varying amount of fluorochrome10. This is in contrast to our method which uses commercially available antibodies and can be adapted to the instrument configuration and can take advantage of the new generation of polymer fluorochromes.
The overall goal of this methodology is to expand the optical limits of most flow cytometers allowing for the recording of five additional markers to interrogate complex cell populations. As a consequence, advanced immunological analysis can be performed on affordable 6-10 fluorochrome flow cytometers, and 2-3 fluorochrome field instruments can achieve remarkable results in areas with limited resources. High-end instruments can also benefit from this approach by using extra fluorochromes to accomplish deeper flow cytometry analysis and to create modular flow cytometric panels targeting several lineages at the same time11. This can potentially reduce the number of panels used in modular immunophenotyping flow cytometry and reduce costs, errors and handling time. This approach is also very well suited in the case of clinical samples with limited number of cells.

<table>
	<tbody>
		<tr>
			<td>CD3</td>
			<td>BD Biosciences</td>
			<td>562426</td>
			<td>Antibody for staining 
			RRID: AB_11152082</td>
		</tr>
		<tr>
			<td>CD56</td>
			<td>BD Biosciences</td>
			<td>562751</td>
			<td>Antibody for staining 
			RRID: AB_2732054</td>
		</tr>
		<tr>
			<td>TCRgd</td>
			<td>BD Biosciences</td>
			<td>331217</td>
			<td>Antibody for staining 
			RRID: AB_2562316</td>
		</tr>
		<tr>
			<td>CD4</td>
			<td>BD Biosciences</td>
			<td>555347</td>
			<td>Antibody for staining 
			RRID: AB_395752</td>
		</tr>
		<tr>
			<td>CD8</td>
			<td>BD Biosciences</td>
			<td>555367</td>
			<td>Antibody for staining 
			RRID: AB_395770</td>
		</tr>
		<tr>
			<td>CD14</td>
			<td>BD Biosciences</td>
			<td>555398</td>
			<td>Antibody for staining 
			RRID: AB_395799</td>
		</tr>
		<tr>
			<td>CD19</td>
			<td>BD Biosciences</td>
			<td>555413</td>
			<td>Antibody for staining 
			RRID: AB_395813</td>
		</tr>
		<tr>
			<td>CD3</td>
			<td>BD Biosciences</td>
			<td>555335</td>
			<td>Antibody for staining 
			RRID: AB_398591</td>
		</tr>
		<tr>
			<td>CD56</td>
			<td>BD Biosciences</td>
			<td>555518</td>
			<td>Antibody for staining 
			RRID: AB_398601</td>
		</tr>
		<tr>
			<td>TCRgd</td>
			<td>BD Biosciences</td>
			<td>331211</td>
			<td>Antibody for staining 
			RRID: AB_1089215</td>
		</tr>
		<tr>
			<td>CCR7</td>
			<td>Biolegend</td>
			<td>353217</td>
			<td>Antibody for staining 
			RRID: AB_10913812</td>
		</tr>
		<tr>
			<td>CD45RA</td>
			<td>BD Biosciences</td>
			<td>560674</td>
			<td>Antibody for staining 
			RRID: AB_1727497</td>
		</tr>
		<tr>
			<td>CCR4</td>
			<td>BD Biosciences</td>
			<td>561034</td>
			<td>Antibody for staining 
			RRID: AB_10563066</td>
		</tr>
		<tr>
			<td>CCR6</td>
			<td>BD Biosciences</td>
			<td>353419</td>
			<td>Antibody for staining 
			RRID: AB_11124539</td>
		</tr>
		<tr>
			<td>CXCR3</td>
			<td>BD Biosciences</td>
			<td>561730</td>
			<td>Antibody for staining 
			RRID: AB_10894207</td>
		</tr>
		<tr>
			<td>CD57</td>
			<td>BD Biosciences</td>
			<td>562488</td>
			<td>Antibody for staining 
			RRID: AB_2737625</td>
		</tr>
		<tr>
			<td>HLA-DR</td>
			<td>BD Biosciences</td>
			<td>563083</td>
			<td>Antibody for staining 
			RRID: AB_2737994</td>
		</tr>
		<tr>
			<td>CD16</td>
			<td>BD Biosciences</td>
			<td>563784</td>
			<td>Antibody for staining 
			RRID: AB_2744293</td>
		</tr>
		<tr>
			<td>Dead cells</td>
			<td>Life technologies</td>
			<td>L-23105</td>
			<td>Live/dead discrimination</td>
		</tr>
		<tr>
			<td>Falcon 5 ml round-bottom polystyrene test tube with cell strainer snap cap</td>
			<td>BD Bioscience</td>
			<td>352235</td>
			<td>to filter cell suspension before passing though the flow cytometer</td>
		</tr>
		<tr>
			<td>Falcon 5 ml round-bottom polystyrene test tube</td>
			<td>BD Bioscience</td>
			<td>352001</td>
			<td>to stain whole blood</td>
		</tr>
		<tr>
			<td>Recovery Cell Culture Freezing Medium</td>
			<td>Thermo fisher</td>
			<td>12648010</td>
			<td>Freezing cells</td>
		</tr>
		<tr>
			<td>96-well V-bottom plate</td>
			<td>Thermo fisher</td>
			<td>249570</td>
			<td>plate for staining</td>
		</tr>
		<tr>
			<td>FACSAria IIu Cell Sorter</td>
			<td>BD Biosciences</td>
			<td></td>
			<td>Flow cytometer</td>
		</tr>
		<tr>
			<td>FCS Express 6</td>
			<td>De Novo Software</td>
			<td></td>
			<td>FACS analysis</td>
		</tr>
		<tr>
			<td>Graphpad Prism</td>
			<td>GraphPad software</td>
			<td></td>
			<td>Data analysis</td>
		</tr>
	</tbody>
</table>

discrimination of seven immune cell subsets by two-fluorochrome flow cytometry

Immune cell characterization heavily relies on multicolor flow cytometry to identify subpopulations based on differential expression of surface markers. Setup of a classic multicolor panel requires high-end instruments, custom labeled antibodies, and careful study design to minimize spectral overlap. We developed a multiparametric analysis to identify major human immune populations (CD4+ and CD8+ T cells, &#947;&#948; T cells, B cells, NK cells and monocytes) in peripheral blood by combining seven lineage markers using only two fluorochromes. Our strategy is based on the observation that lineage markers are constantly expressed in a unique combination by each cell population. Combining this information with a careful titration of the antibodies allows investigators to record five additional markers, expanding the optical limit of most flow cytometers. Head-to-head comparison demonstrated that the vast majority of immune cell populations in peripheral blood can be characterized with comparable accuracy between our method and the classic &#34;one fluorochrome-one marker approach&#34;, although the latter is still more precise for identifying populations such as NKT cells and &#947;&#948; T cells. Combining seven markers using two fluorochromes allows for the analysis of complex immune cell populations and clinical samples on affordable 6-10 fluorochrome flow cytometers, and even on 2-3 fluorochrome field instruments in areas with limited resources. High-end instruments can also benefit from this approach by using extra fluorochromes to accomplish deeper flow cytometry analysis. This approach is also very well suited for screening several cell populations in the case of clinical samples with limited number of cells.

Setup and analysis of a flow cytometry experiment of human peripheral blood cells stained with seven lineage markers (anti-CD3, -CD4, -CD8, -CD14, -CD19, -CD56 and -TCR &#947;&#948; antibodies) using only two fluorochromes are presented.
Representative results are described for anti-CD8 and -CD56 antibody titration. For each antibody (in this example, anti-CD8), data from ten successive 2-fold dilutions were recorded to calculat...

Watch this Scientific Journal Video about Discrimination of Seven Immune Cell Subsets by Two-fluorochrome Flow Cytometry at JoVE.com

Discrimination of Seven Immune Cell Subsets by Two-fluorochrome Flow Cytometry

Here, we present a flow cytometric protocol to identify CD4+ and CD8+ T cells, &#947;&#948; T cells, B cells, NK cells and monocytes in human peripheral blood by using only two fluorochromes instead of seven. With this approach, five additional markers can be recorded on most flow cytometers.

Immune cell characterization heavily relies on multicolor flow cytometry to identify subpopulations based on differential expression of surface markers. ...

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Research

JoVE Journal

Immunology and Infection

13.7K Views.  Medimmune, LLC. Here, we present a flow cytometric protocol to identify CD4+ and CD8+ T cells, γδ T cells, B cells, NK cells and monocytes in human peripheral blood by using only two fluorochromes instead of seven. With this approach, five additional markers can be recorded on most flow cytometers.

Video: Discrimination of Seven Immune Cell Subsets by Two-fluorochrome Flow Cytometry

Multicolor Flow Cytometry Analyses of Cellular Immune Response in Rhesus Macaques

The rhesus macaque model is currently the best available model for HIV-AIDS with respect to understanding the pathogenesis as well as for the development of vaccines and therapeutics1,2,3. Here, we describe a method for the detailed phenotypic and functional analyses of cellular immune responses, specifically intracellular cytokine production by CD4+ and CD8+ T cells as well as the individual memory subsets. We obtained precise quantitative and qualitative measures for the production of interferon gamma (INF-) and interleukin (IL) -2 in both CD4+ and CD8+ T cells from the rhesus macaque PBMC stimulated with PMA plus ionomycin (PMA+I). The cytokine profiles were different in the different subsets of memory cells. Furthermore, this protocol provided us the sensitivity to demonstrate even minor fractions of antigen specific CD4+ and CD8+ T cell subsets within the PBMC samples from rhesus macaques immunized with an HIV envelope peptide cocktail vaccine developed in our laboratory. The multicolor flow cytometry technique is a powerful tool to precisely identify different populations of T cells 4,5 with cytokine-producing capability6 following non-specific or antigen-specific stimulation 5,7.

We demonstrate the utility of multicolor flow cytometry for detailed phenotypic and functional characterization of total as well as memory subsets of CD4+ and CD8+ T cells in rhesus macaques, the ideal model for HIV/AIDS vaccine studies.

The rhesus macaque model is currently the best available model for HIV-AIDS with respect to understanding the pathogenesis as well as for the development ...

Measuring Calpain Activity in Fixed and Living Cells by Flow Cytometry

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, including signaling, cytoskeletal remodeling, regulation of gene expression, apoptosis and cell cycle progression 1. Calpains have been implicated in many pathologies including muscular dystrophies, cancer, diabetes, Alzheimer's disease and multiple sclerosis 1. Calpain regulation is complex and incompletely understood. mRNA and protein levels correlate poorly with activity, limiting the use of gene or protein expression techniques to measure calpain activity. This video protocol details a flow cytometric assay developed in our laboratory for measuring calpain activity in fixed and living cells. This method uses the fluorescent substrate BOC-LM-CMAC, which is cleaved specifically by calpain, to measure calpain activity. 2 In this video, calpain activity in fixed and living murine 32Dkit leukemia cells, alone or as part of a splenocyte population is measured using an LSRII (BD Bioscience). 32Dkit cells are shown to have elevated activity compared to normal splenocytes.

This article will detail the protocol for measuring calpain activity in fixed and living cells using flow cytometry.

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, ...

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.

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

Seven Steps to Stellate Cells

Hepatic stellate cells are liver-resident cells of star-like morphology and are located in the space of Disse between liver sinusoidal endothelial cells and hepatocytes1,2. Stellate cells are derived from bone marrow precursors and store up to 80% of the total body vitamin A1, 2. Upon activation, stellate cells differentiate into myofibroblasts to produce extracellular matrix, thus contributing to liver fibrosis3. Based on their ability to contract, myofibroblastic stellate cells can regulate the vascular tone associated with portal hypertension4. Recently, we demonstrated that hepatic stellate cells are potent antigen presenting cells and can activate NKT cells as well as conventional T lymphocytes5. 

 Here we present a method for the efficient preparation of hepatic stellate cells from mouse liver. Due to their perisinusoidal localization, the isolation of hepatic stellate cells is a multi-step process. In order to render stellate cells accessible to isolation from the space of Disse, mouse livers are perfused in situ with the digestive enzymes Pronase E and Collagenase P. Following perfusion, the liver tissue is subjected to additional enzymatic treatment with Pronase E and Collagenase P in vitro. Subsequently, the method takes advantage of the massive amount of vitamin A-storing lipid droplets in hepatic stellate cells. This feature allows the separation of stellate cells from other hepatic cell types by centrifugation on an 8% Nycodenz gradient. The protocol described here yields a highly pure and homogenous population of stellate cells. Purity of preparations can be assessed by staining for the marker molecule glial fibrillary acidic protein (GFAP), prior to analysis by fluorescence microscopy or flow cytometry. Further, light microscopy reveals the unique appearance of star-shaped hepatic stellate cells that harbor high amounts of lipid droplets. 


 Taken together, we present a detailed protocol for the efficient isolation of hepatic stellate cells, including representative images of their morphological appearance and GFAP expression that help to define the stellate cell entity.

Here we describe a method for the isolation of hepatic stellate cells from mouse liver. For stellate cell purification, mouse livers are digested in situ and in vitro by pronase-collagenase treatment prior to density gradient centrifugation. This technique yields highly pure hepatic stellate cells.

Hepatic stellate cells are liver-resident  cells of star-like morphology and are located in the space of Disse between  liver sinusoidal endothelial cells ...

Flow Cytometry Analysis of Immune Cells Within Murine Aortas

Atherosclerosis is a chronic inflammatory process of medium and large size vessels that is characterized by the formation of plaques consisting of foam cells, immune cells, vascular endothelial and smooth muscle cells, platelets, extracellular matrix, and a lipid-rich core with extensive necrosis and fibrosis of surrounding tissues.1 The innate and adaptive arms of the immune response are involved in the initiation, development and persistence of atherosclerosis.2, 3 There is a significant body of evidence that different subsets of the immune cells, such as macrophages, dendritic cells, T and B lymphocytes, are present within the aortas of healthy and atherosclerosis-prone mice4. Additionally, immune cells are found in the surrounding aortic adventitia which suggests an important role of this tissue in atherogenesis.2
For some time, the quantitative detection of different types of immune cells, their activation status, and the cellular composition within the aortic wall was limited by RT-PCR and immunohistochemical methods for the study of atherosclerosis. Few attempts were made to perform flow cytometry using human aortas, and a number of problems, such as a high autofluorescence, have been reported5,6. Human atherosclerotic plaques were digested with collagenase 1, and free cells were collected and stained for CD14+/CD11c+ to highlight macrophage-derived foam cells. In this study, a "mock" channel was used to avoid false-positive staining.6 Necrotic materials accumulating during the digestion process give rise in a large amount of debris that generates a high autofluorescence in aortic samples. To resolve this problem, a panel of negative and positive controls has been proposed, but only double staining could be applied in these samples. We have developed a new flow cytometry-based method7 to analyze the immune cell composition and characterize the activation, proliferation, differentiation of immune cells in healthy and atherosclerosis-prone aorta. This method allows the investigation of the immune cell composition of the aortic wall and opens possibilities to use a broad spectrum of immunological methods for investigations of immune aspects of this disease.

This paper presents a flow cytometry-based method to investigate the immune composition of aortas. The paper also illustrates an additional technique that allows examining surrounding adventitia and vessel wall separately. This method opens possibilities to perform phenotypical analyses of aortic leukocytes and apply several immunological assays for atherosclerosis studies.

Atherosclerosis is a chronic inflammatory process of medium and large size vessels that is characterized by the formation of plaques consisting of foam ...

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.

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

Detection of Fluorescent Nanoparticle Interactions with Primary Immune Cell Subpopulations by Flow Cytometry

Engineered nanoparticles are endowed with very promising properties for therapeutic and diagnostic purposes. This work describes a fast and reliable method of analysis by flow cytometry to study nanoparticle interaction with immune cells. Primary immune cells can be easily purified from human or mouse tissues by antibody-mediated magnetic isolation. In the first instance, the different cell populations running in a flow cytometer can be distinguished by the forward-scattered light (FSC), which is proportional to cell size, and the side-scattered light (SSC), related to cell internal complexity. Furthermore, fluorescently labeled antibodies against specific cell surface receptors permit the identification of several subpopulations within the same sample. Often, all these features vary when cells are boosted by external stimuli that change their physiological and morphological state. Here, 50 nm FITC-SiO2 nanoparticles are used as a model to identify the internalization of nanostructured materials in human blood immune cells. The cell fluorescence and side-scattered light increase after incubation with nanoparticles allowed us to define time and concentration dependence of nanoparticle-cell interaction. Moreover, such protocol can be extended to investigate Rhodamine-SiO2 nanoparticle interaction with primary microglia, the central nervous system resident immune cells, isolated from mutant mice that specifically express the Green Fluorescent Protein (GFP) in the monocyte/macrophage lineage. Finally, flow cytometry data related to nanoparticle internalization into the cells have been confirmed by confocal microscopy.

Analysis of nanoparticle interaction with defined subpopulations of immune cells by flow cytometry.

Engineered nanoparticles are endowed with very promising properties for therapeutic and diagnostic purposes. This work describes a fast and reliable ...

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

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 ...

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.

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

Discrimination of Seven Immune Cell Subsets by Two-fluorochrome Flow Cytometry

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

Measurement of T Cell Alloreactivity Using Imaging Flow Cytometry

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages