An In Vitro Assay for Studying the Cytotoxicity of Pre-Activated CD8+ T Cells against Cancer Cells

Take two CD8⁺ T cell populations: effector  CD8⁺ T cells pre-activated through antibody-mediated engagement of T cell and co-stimulatory receptors and suppressed CD8⁺ T cells inhibited by myeloid-derived suppressor cells.

Add these T cells to basement-membrane-coated wells with cancer cells expressing a nuclear-restricted fluorescent protein.

Pipette a fluorogenic  caspase-3 substrate — a DNA-binding fluorogenic dye conjugated to a caspase-3 recognition site-containing peptide.

Incubate. Effector T cell receptors bind to the class I MHC-peptide complex on the cancer cells, triggering the release of cytotoxic granules containing perforins and granzymes.

Perforins create cancer cell membrane pores, allowing granzyme entry and caspase activation, initiating apoptosis.

Activated caspases additionally cleave the fluorogenic caspase substrate, releasing the dye that binds to DNA and fluoresces.

Under a fluorescence microscope, cancer cells co-cultured with effector T cells exhibit dual nuclear fluorescence, indicating cancer cell apoptosis.

In contrast, cancer cells co-cultured with suppressed T cells display only nuclear protein fluorescence, suggesting non-apoptotic cells.

To set up a pre-activated CD8-positive T cell target cell co-culture, first, add 30 microliters of 1 to 100 diluted growth-factor reduced soluble basement membrane matrix to the appropriate wells of a 96-well flat-bottom plate suitable for microscopy. Shake the plate to spread the matrix evenly, and place the plate in the cell culture incubator for at least one hour. While the plate is equilibrating, prepare the target cells. Aspirate the medium, wash with PBS, and add 1 milliliter of 0.05% trypsin-EDTA at room temperature for 1 minute.

Add 9 milliliters of DMEM supplemented with fetal bovine serum by gentle pipetting, and transfer the dissociated cells into tubes. After centrifuging the cells, resuspend the pellet in 500 microliters of EDMEM. Filter the resuspended cells through a cell strainer, and count the live cells. Then, adjust the density to 4 times 10 to the fourth target cells per milliliter in fresh EDMEM on ice.

Next, pipette the pre-activated CD8-positive T cells a few times to resuspend the cells into a single-cell suspension, and transfer the floating T cells into one 1.5-milliliter tube per condition. Collect any remaining cells with 200 microliters of PBS per well, transfer the washes into the appropriate 1.5-milliliter tubes, and centrifuge. Aspirate the medium, and add 1 milliliter of EDMEM to each tube for centrifugation.

After centrifugation, resuspend the pellets in 100 microliters of fresh EDMEM per tube. After counting, adjust the cells in each tube to a density of 1.6 times 10 to the fifth pre-activated CD8-positive T cells per milliliter of medium, and place the cells on ice. When the cells are ready, aspirate the basement membrane matrix from each well of the 96-well microscopy plate, and add 50 microliters of target cells to individual wells within the inner 60 wells of the plate with mixing.

Add 25 microliters of EDMEM supplemented with 4 times 10 to the third units per milliliter of IL-2, and 10 micromolar fluorogenic caspase-3 substrate to the appropriate wells. Then, add 25 microliters of pre-activated CD8-positive T cells to the appropriate wells, and finally, add EDMEM to the appropriate wells to make a total volume of up to 100 microliters. Add 200 microliters of PBS or sterile water into all of the empty wells, shake the plate, and leave the plate on a flat surface for 10 minutes.

To image the cells, set the microscope to acquire images in phase contrast, as well as fluorescent channels suitable for the nuclear-restricted fluorescent protein, and the fluorogenic activated caspase-3 substrate fluorophores used in the experiment. Then, capture images of each experimental well in phase contrast, and the two fluorescent channels every one to three hours for at least 72 hours.