The overall goal of this methodology is to expand beyond the optical limit of most flow cytometers by allowing a researcher to record five additional markers to interrogate complex cell populations. Using this technique, it is possible to achieve a deep immunophenotyping when working with instrument with a low number of detectors or sample with limited number of cells. Begin this procedure by carefully transferring drawn blood into a 50-milliliter conical tube.
Dilute the blood with an equal amount of PBS without calcium and magnesium. Add 15 milliliters of density gradient medium to the bottom of a new 50-milliliter conical tube. Carefully overlay the diluted blood on top of the density gradient medium, avoiding any mixing between the density gradient medium and diluted blood.
Centrifuge at 400 times g for 30 minutes at room temperature with no brake to avoid disruption of the interface. After centrifugation, use a pipette to carefully aspirate and discard the upper layer, paying attention to not remove cells at the interface between the plasma and density gradient medium. Collect as many peripheral blood mononuclear cells, or PBMCs, as possible from the interface without touching the red cell pellet at the bottom of the 50-milliliter conical tube, and transfer to a new tube.
Add PBS to bring the final volume to 25 milliliters, and invert several times to mix. Then, wash the PBMCs twice as indicated in the text. After removing the platelets and counting the PBMCs as described in the text, resuspend the cells in PBS, 0.1%sodium azide to a concentration of one times 10 to the seventh cells per milliliter.
To prepare the PBMCs for staining, transfer 100 microliters of each sample to a 96-well V-bottom plate. Centrifuge the plate at 350 times g for three minutes at room temperature. Carefully aspirate the supernatant without disturbing the cell pellet.
To each well, add 100 microliters of PBS containing a live/dead fixable dye that reacts with free amine on proteins, and resuspend the PBMC mixture carefully. Subsequently, allow the plate to sit for 10 minutes at room temperature for labeling of dead cells. For each sample, prepare 30 microliters of a mix containing all seven antibodies.
At this stage, titrated antibodies against different target molecules and in different fluorochromes can be added as well. Centrifuge the 96-well plate at 350 times g for three minutes at room temperature, 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 minutes at room temperature in the dark. After 30 minutes, add 150 microliters of staining buffer to each well, and centrifuge the plate at 350 times g for three minutes at room temperature. Carefully aspirate the supernatant without disturbing the cell pellet, and resuspend the cells in 200 microliters of PBS.
The cells are now ready for flow cytometry analysis. Anti-CD3, CD8, CD14, CD19, and TCR gamma delta antibodies are titrated with a maximum staining index curve. Antibody titration is the most critical step in this protocol to properly identify seven immune cell subset using two fluorochromes.
To prepare a two-fold antibody dilution, fill 10 wells of a 96-well plate with 40 microliters of staining buffer per well. For the purposes of this video, only the titration of anti-CD8 will be shown. In the first well, increase the final volume to 80 microliters of staining buffer, and add anti-CD8 at a concentration four times the concentration suggested by the manufacturer.
Mix well and transfer 40 microliters to the second well. Mix well and repeat this step for all the other wells. Stain 10 samples of PBMCs or whole blood with 30 microliters of the 10 different two-fold dilutions of anti-CD8 following the protocol demonstrated earlier.
Acquire data with a flow cytometer, and plot the signal from each dilution. After calculating the stain index for each antibody concentration as detailed in the text, plot the stain index versus the antibody concentration expressed as a fraction of the antibody dilution, and identify the concentration of antibody with the maximum stain index value. To titrate the anti-CD4 and anti-CD56 antibodies, start with the two-fold dilution strategy demonstrated earlier, adding additional concentrations in between to finely identify the range of concentration that will allow separation of CD4-positive T cells and NK cells from the other cell populations.
Titrate the anti-CD4 antibody by placing the anti-CD4 signal between the double CD8-positive, CD3-positive signal and the CD3 single positive population. 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. To begin this two-fluorochrome, seven-marker gating strategy, select the lymphocyte gate, and create a dot plot with one of the two fluorochromes used in this protocol on each axis.
Gate on CD8-positive T cells identified as CD3 and CD8 double positive cells at the top right corner of the dot plot. Exclude the dim CD8 population which might contain NKT cells. Gate on CD4-positive T cells identified as the population in between CD8-positive T cells and the CD3 single positive populations.
Gate on gamma delta T cells identified as high CD3 cells. Subdivide gamma delta T cells to CD8 positive and CD8 negative. Gate on NK cells identified as the population in between CD3-positive and CD3-negative cells.
Subdivide the NK cells to CD8 positive and CD8 negative. Gate on B cells identified as the CD3-negative, CD19-positive population on the lower right corner of the dot plot. Select the monocyte gate, and create a dot plot with one of the two fluorochromes used in this protocol on each axis.
Gate on the CD3-negative, CD14-positive population. Using this strategy, lymphocytes from a patient with multiple myeloma were gated on the basis of their forward scatter and side scatter and their flow cytometric profile. CD8-positive memory subpopulations and naive T cells were identified by the expression of CD45RA and CCR7.
Expression of HLA-DR and CD57 was used to study T cell activation in CD8-positive naive, memory, central memory, effector memory, and effector memory CD45RA-positive cells. In a similar manner, CD4-positive naive and memory T cells were identified. Within the memory population, CCR4 and CCR6 were used to identify T helper subpopulations in CD4-positive T cells.
Subpopulations of NK cells were characterized by CD16 and CD57 expression. This strategy was used to investigate the dynamics in immune populations of longitudinal samples from a donor with multiple myeloma receiving a stem cell transplant. Characterization of CD8 and CD4 subpopulations over time showed a sustained CD4-positive and CD8-positive T cell activation and a skewing of T helper to a T-helper-one phenotype.
But at day 60, the percentage of B cells dramatically augmented, predicting the patient relapse. Proper cell preparation and antibody titration are critical steps to achieve reliable results using this technique. Antibody targeting other markers can be added to accomplish deeper flow cytometry analysis and to create modular flow cytometric panels targeting several lineages at the same time.
Similar approaches aimed at expanding the number of recordable markers could also be developed using different sets of marker or for use in different animal models. Do not forget that sodium azide can cause burns to skin and eyes. Therefore, a lab coat, protective glasses, and gloves should always be worn when handling this reagent.
