This article describes a method for characterizing CD8 + memory T cell subsets in non-human primates using flow cytometry. The protocol involves the use of pMHC-I tetramers and various antibodies to identify and analyze T cell populations.
Take non-human primate peripheral blood mononuclear cells, or PBMCs, containing CD8+ memory T cells against simian immunodeficiency virus or SIV.
Add a protein kinase inhibitor to prevent internalization of the T cell receptor, or TCR upon subsequent stimulation.
Introduce peptide-major histocompatibility complex class I, or pMHC-I tetramers, fluorophore-conjugated complexes comprising four MHC-I molecules bound to SIV-specific peptides. These peptides bind to TCRs on the surface of T cells.
Add a mixture containing fluorophore-conjugated antibodies binding to CD8, CD28, and CCR7 — markers delineating memory T cell subsets — and antibodies labeling non-CD8+ cells.
A fluorescent viability dye labels dead cells.
Fix the cells. Permeabilize them to access intracellular targets.
Add fluorophore-conjugated antibodies binding CD3, a T cell marker, and intracellular granzyme B, an activated CD8+ T cell marker.
Using flow cytometry, exclude non-CD8+ T cells and dead cells.
Gate pMHC-I tetramer positive CD8+ T cells to characterize different memory CD8+ T cell subsets based on granzyme B, CD28, and CCR7 levels.
Begin by resuspending the Rhesus PBMCs in R-10 medium at a 1.6 times 10 to the seventh cells per milliliter concentration. Aliquot 50 microliters of cells to the appropriate number of flow cytometry tubes, and add 50 microliters of protein kinase inhibitor to each tube. Vortex each tube, and incubate the samples at 37 degree Celsius for 30 minutes.
The correct amount of titrated peptide MHC tetramer must be diluted enough master mix to stain all of the experimental tubes. Therefore, it is important to prepare a 15% excess of the master mix to account for pipetting errors.
While the cells are incubating, pellet the protein aggregates and the tetramer solution by centrifugation to minimize the background staining. Next, dilute the tetramers in enough stain buffer so that 25 microliters of the tetramer master mix can be added to each flow cytometry tube. Then, vortex the sample tubes, and incubate the PBMC tetramer solutions in the dark at room temperature for 45 minutes. At the end of the incubation, add 50 microliters of staining mastermix to the appropriate sample tubes, and mix the tubes by vortexing.
After 25 minutes in the dark at room temperature, wash the cells with wash buffer, pellet the cells by centrifugation, and carefully decant the supernatant without disturbing the pellets. Vortex the cells in the leftover buffer in each tube, and fix the samples with 250 microliters of 2% paraformaldehyde per tube. Immediately vortex and incubate the tubes in the dark at 4 degrees Celsius. After 20 minutes, wash the cells in fresh wash buffer, pellet the cells by centrifugation, and decant the supernatant without disturbing the pellet.
Resuspend the cells in 500 microliters of permeabilization buffer, vortex, and incubate at room temperature for 10 minutes in the dark. Then, wash and pellet the cells. Then, decant the supernatants, as described previously. Proceed to adding 50 microliters of intracellular stain master mix to the appropriate tubes. Vortex each sample and incubate at room temperature in the dark for 30 minutes. After the incubation is done, wash and pellet the cells. The samples are now ready to be acquired in a flow cytometer.
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