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

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

1. Isolation of peripheral blood cells (PBMC) from whole blood
   1. 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.
   2. 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.
   3. 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.
   4. Centrifuge at 400 x g for 30 min at room temperature (RT), with no brake to avoid disruption of the interface.
   5. 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.
   6. 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.
   7. 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.
   8. 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.
      ​CAUTION: Sodium azide can cause death by affecting the central nervous system. Contact may cause burns to skin and eyes.
   9. Dilute the cells for counting by transferring 30 µL of PBMCs in a 1.5 mL microcentrifuge tube. Then add 120 µL of PBS and 150 µL of Trypan blue to determine cell number and viability (1:10 cell dilution). Resuspend carefully.
   10. Transfer 10 µL 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.
   11. 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.
2. 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.
   ​CAUTION: DMSO can be slightly hazardous in case of inhalation (lung irritant), skin contact (irritant, permeator), of eye contact (irritant), of ingestion.
   1. Remove frozen PBMC from liquid nitrogen and place on ice. Transfer the cryovial from the ice directly into the 37 °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.
   2. Remove any water with a 70% ethanol sprayed wipe. Slowly add 1 mL of PBS/0.1% sodium azide at 4 °C and carefully transfer the cell to a 15 mL conical tube. Slowly add cold PBS/0.1% sodium azide at 4 °C to reach a final volume of 15 mL.
   3. 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.
   4. Dilute the cell for counting by transferring 30 µL of PBMCs in a 1.5 mL microcentrifuge tube. Then add 120 µL of PBS and 150 µL of Trypan blue to determine cell number and viability (1:10 cell dilution). Resuspend carefully.
   5. Transfer 10 µL 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.
   6. Resuspend the cells at 10 x 10⁶ per mL in PBS/0.1% sodium azide.
3. Preparation of cells from whole blood
   1. 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.
   2. Transfer 200 µL of whole blood to a 12 mm x 75 mm capped tube, add 2 µL of sodium azide and vortex gently for 2 s.

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.

1. Staining of fresh and frozen PBMC
   1. Transfer 100 µL 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.
   2. 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 µL of PBS containing a live/dead fixable dye that reacts with free amine on proteins for 10 min to label dead cells.
   3. Prepare for each sample 30 µL of a mix containing all the antibodies (anti-CD3. -CD56, TCRγδ 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 µL to easily integrate this methodology to already existing staining protocols.
   4. 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 °C with similar results.
   5. Add 150 µL 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 µL 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 °C overnight and then acquired on a flow cytometer the following day.
      CAUTION: Paraformaldehyde is harmful if swallowed and can cause skin irritation
2. Staining of whole blood
   1. To each 12 mm x 75 mm capped tube, add the cocktail of antibodies (anti-CD3. -CD56, TCRγδ 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 °C with similar results.
   2. 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’s instructions.
      NOTE: The lysis solution should be at RT before use.
   3. 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.
   4. Spin down at 300 x g for 5 min. Carefully aspirate the supernatant without disturbing the cell pellet.
   5. 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 µL of PBS.
   6. Strain the cells through 12 mm x 75 mm tubes with 40 µm filter caps to remove cell aggregates and acquire data on a flow cytometer.

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 γδ follows the standard procedure by which the concentration of antibody to optimally separate positive and negative peaks is derived by maximum staining index^13,14. 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+ γδ 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.

1. Maximum stain index curve
   NOTE: Titration of anti-CD3, -CD8, -CD14, -CD19 and -TCR γδ 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.
   1. Prepare a 2-fold antibody dilution by filling 10 wells of a 96-well plate with 40 µL of staining buffer. In the first well, increase the final volume to 80 µL of staining buffer and add the antibody of interest at a concentration 4 times the concentration suggested by the manufacturer.
   2. Mix well and transfer 40 µL to the second well. Mix well and repeat this step for the all the other wells.
   3. Stain 10 samples of PBMC or whole blood with 30 µL of the 10 different 2-fold dilutions of antibodies following the protocol described before.
   4. Acquire data with a flow cytometer and plot the signal from each dilution (Figure 1A).
   5. 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.
   6. 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 — median fluorescent intensity of the negative population) ÷ (2 x standard deviation of the fluorescent intensity of the negative population) (Figure 1B).
   7. 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).
2. 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).
   1. 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⁺ T cells and NK cells from the other cell populations.
   2. 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.
   3. 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.

4. Gating Strategy

1. Identify lymphocytic and monocytic cell populations and remove dead cells and most of the residual red blood cells from the analysis.
   1. 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).
   2. 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.
2. Two-fluorochrome seven-marker gating strategy of the lymphocytic populations.
   NOTE: Within the CD3 positive subgroup, CD4⁺, CD8⁺ and γδ T cells can be separated using antibodies that solely target CD4, CD8 and the γδ 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.
   1. Select the lymphocyte gate and create a dot plot with on each axis one of two fluorochromes used in this protocol (Figure 2B).
   2. 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 γδ T cells identified as high CD3 cells. Subdivide γδ T cells in CD8 positive and CD8 negative.
   3. 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.
   4. 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.