This assay can be employed to quantify T-cell-mediated cytotoxicity against two distinct peptide epitopes of tumor antigens simultaneously. The in vivo nature of this assay allows for cytotoxic function of T-cells to be measured within the intact architecture of a secondary lymphoid organ. In addition, the killing observed is an accurate reflection of the absolute number of T-cells, and not just their frequencies, which can be misleading.
This assay enables the examination of cytotoxic T cell responses to immunodominant and sub-dominant tumor antigens. It therefore provides valuable information when you're designing a vaccine, as well as immunotherapeutic protocols for cancer. Our protocol has been tailored to study CD8 T cell responses.
However, it can be modified to measure cytotoxicity elicited by other killer cell types, such as NK cells and innate-like T lymphocytes. This assay requires experience in aseptic technique, mouse tissue handling, target cell prep, as well as expertise in tail vein injections. One needs to acquire this skill set before attempting the in vivo killing assay.
Once the T-antigen-positive adherent tumor cell line has become fully confluent or slightly over-confluent, gently rinse the monolayer with sterile, warm PBS, and detach the tumor cells with 0.25%trypsin EDTA in a biosafety cabinet. Tap the sides of the culture flask several times to release the remaining adherent cells, and stop the reaction after about five minutes with the addition of five milliliters of DMEM medium. Pipette the cell solution a few times before filtering the cell suspension through a 70-micrometer-pore cell strainer into a 50-milliliter conical tube.
Collect the cells by centrifugation, and resuspend the pellet in 10 milliliters of sterile, cold PBS for the first of three washes under the same conditions. After the third wash, resuspend the cells at a four times 10 to the seven cells per milliliter of sterile PBS concentration, and inject 500 microliters of the T-antigen-positive tumor cell suspension intraperitoneally into each six-to-12-week-old C57 Black 6 mouse recipient. For target splenocyte preparation, place the euthanized donor mouse in the prone position, and spray the animal with 70%ethanol.
Using sterile forceps and scissors, lift the skin and make a small ventral midline incision. Cut the skin in a cross-like fashion to expose the peritoneum, and use the forceps to pull up the peritoneum in a tent-like fashion, without catching any of the internal organs. Cut open the peritoneum to expose the peritoneal cavity, and gently transfer the spleen to a 15-milliliter Dounce tissue grinder containing five milliliters of sterile PBS.
Use the plunger to apply manual pressure until the splenic tissue dissipates into a red homogeneous cell suspension, and transfer the homogenate into a 15-milliliter tube. Collect the suspension by centrifugation, and resuspend the pellet in four milliliters of ammonium chloride potassium lysine buffer to eliminate the erythrocytes. After four minutes, stop the process with eight milliliters of complete medium, and filter the cell suspension through a 70-micrometer-pore cell strainer into a new 50-milliliter tube.
Separate the white blood cells from the red blood cell debris by centrifugation, and resuspend the pellet in 12 milliliters of complete medium. Then split the splenocyte suspension into three equal four-milliliter aliquots. For peptide coding of the target splenocytes, pulse one splenocyte aliquot with one micromolar of irrelevant peptide, one aliquot with one micromolar of Site I peptide, and one aliquot with one micromolar of Site IV peptide for one hour at 37 degrees Celsius and 5%carbon dioxide.
At the end of the incubation, remove the excess peptide by centrifugation and wash the peptide-pulsed splenocytes two times in 12 milliliters of sterile, cold PBS per wash per tube. After the second wash, resuspend the cells in four milliliters of fresh, sterile PBS per tube. Add 025, 0.25, and two-micromolar CFSE into the irrelevant, Site I, and Site IV peptide-pulsed splenocyte suspensions respectively.
Place the tubes at 37 degrees Celsius for 15 minutes with manual inversion once every five minutes. At the end of the incubation, add three milliliters of heat-inactivated fetal bovine serum to each tube to stop the CFSE reaction and bring the final volume in each tube up to 15 milliliters with PBS. Then collect the dye-labeled cells by centrifugation for two washes in 12 milliliters of fresh, sterile PBS per wash.
To confirm an adequate CFSE labeling of the target splenocytes, resuspend the pellet in three milliliters of PBS and mix with gentle vortexing. Transfer 10 microliters from each CFSE-labeled cell suspension into an individual five-milliliter round bottom polystyrene fluorescence-activated cell sorting, or FACS tube, containing PBS, and load the tube onto a flow cytometer equipped with a 488-nanometer laser. In the flow cytometer software, create a lymphocyte gate based on the forward scatter and side scatter properties of the cells before acquiring 5, 000 events falling within the lymphocyte gate in the Fluorescence 1 channel.
Within the parent CFSE-positive population, draw additional histogram gates to identify the CFSE-low, CFSE-intermediate, and CFSE-high subpopulations. Then confirm an equal or a near-equal event number within the three gates for the other two tubes of labeled cells. For injection of the target cells, first gently vortex the source tubes and pool the three CFSE-labeled cell suspensions into a new tube at equal ratios.
Top up the tube contents with sterile PBS, and collect the cells by centrifugation for counting. Then adjust the volume to a one times 10 to the seven mixed target cells per 200 microliters of PBS per recipient concentration, and inject 200 microliter volumes of the cell suspension into the tail vein of each recipient C57 Black 6 mouse. Two or four hours after the injection, remove and process the spleen of each recipient animal as demonstrated, and resuspend the isolated white blood cells in three milliliters of PBS per spleen.
Transfer approximately one times 10 to the seven cells from each processed spleen into a clean FACS tube, and immediately run the cells on the flow cytometer as demonstrated to gate the CFSE-low, intermediate, and high target cell populations. Then acquire a total of 2, 000 CFSE-low events in the Fluorescence 1 channel, and calculate the specific lysis of each cognate target cell population according to the formula detailed in the text. In this representative experiment, the success of the depletion of naturally occurring regulatory T-cells in mice injected with anti-CD25 monoclonal antibody was confirmed by flow cytometry.
As expected, near-equal peaks corresponding to the control and cognate target cells were detectable in naive mice. In contrast, Site-IV-displaying target cells were almost completely absent in T-antigen-primed mice, regardless of their prior treatment with anti-CD25 monoclonal antibody or PBS. Interestingly, naturally occurring T reg cell depletion by anti-CD25 monoclonal antibody augmented in vivo cytotoxic T-lymphocyte-mediated lysis of Site-I-pulsed target cells.
In this second representative investigation, programmed death 1 blockade enhanced sub-dominant CD8-positive T cell sites one and two-slash-three-specific responses, suggesting that interfering with programmed death 1/programmed death ligand 1 interactions may induce epitope-spreading in anti-cancer CD8-positive T cell responses. It's critical when labeling your donor cells with CFSE, make sure your concentrations are precise. Otherwise this may result in overlapping histogram peaks, which make your required calculations and data interpretation difficult, if not impossible.
It will be important to optimize in vivo cytotoxicity assays to quantify effector functions elicited by different killer cell types that recognize peptide and non-peptide antigens in the same host.
