A Real-Time In Vitro Cytolysis Assay to Evaluate the Potency of CAR T Cells

1. Generation of CAR-encoding lentivirus

NOTE: Once the specific chimeric antigen receptor (CAR) T-cell plasmid construction is completed (CD47 and others), lentiviral CARs are generated by the standard procedure, using 293 FT cells, a lentiviral packaging mix, and transfection agents (see the Table of Materials) as described. Subsequently, use a quantitative reverse transcription polymerase chain reaction (RT-PCR) kit and a thermal cycler (see  Table of Materials) to determine the virus titer by measuring the lentiviral RNA amount according to the manufacturer's protocol. It is important that all lentiviral procedures be carried out strictly following safety requirements.

1. Seed 15 x 10⁶ HEK293FT cells in Dulbecco's modified Eagle's medium (DMEM), and incubate the cells overnight at 37 °C in one 150 mm dish inside a humidified 5% carbon dioxide (CO₂) incubator.
2. Prepare two 15 mL tubes with transfection complex. The first tube contains lentiviral vector plasmid DNA (5 µg) and lentiviral packaging mix (22.5 µg) in 2.5 mL of transfection dilution solution. The second tube contains 82.5 µL of transfection reagent in 2.5 mL of transfection dilution solution (see the Table of Materials).
3. Pipet the contents of tube 1 into tube 2, and incubate the mixture at room temperature for 15 min.
4. Transfer the contents of the tube dropwise to the dish of HEK293FT cells and incubate the sample overnight at 37 °C in a humidified 5% carbon dioxide, CO₂ incubator.
5. The next day, replace the existing medium with 19 mL of fresh DMEM culture medium and continue to incubate the cells overnight inside the humidified 5% CO₂ incubator at 37 °C.
6. Transfer the medium from the dish to a 50 mL centrifuge tube. Keep the tube with the virus-containing medium in the refrigerator.
7. Repeat the above procedure, adding fresh DMEM and collecting it again after 1 day.
8. Combine two collections of the media into one centrifuge tube. Centrifugate the tube at 2,000 x g for 30 min at 4 °C.
9. Transfer most of the lentivirus-containing supernatant to an ultraclear centrifuge tube. Leave a minimum volume, about 1 mL of the supernatant to avoid disturbing the pellet which may contain cells and/or debris.
10. Ultracentrifuge the above-clarified supernatant at 110,000 x g for 100 min at 4 °C.
11. Remove the supernatant carefully and gently add 100 µL of DMEM medium to the virus pellet at the bottom of the tube. Leave the tube on ice for 15 min. Mix the solution gently and aliquot the lentivirus solution into prechilled sterile tubes. Store these virus stock tubes in a -80 °C freezer.
12. Use a quantitative RT-PCR kit to determine the titer of the lentivirus according to the manufacturer's protocol, which extracts and measures lentiviral RNA.

2. Generation and expansion of CAR T cells

1. Activate previously frozen human peripheral blood mononuclear cells (PBMCs) (about 1 x 10⁶ to 2 x 10⁶ cells) in 1 mL of CAR T-cell medium with an equal number of CD3/CD28-coated microbeads (see the Table of Materials) and incubate the cells at 37 °C in a humidified 5% CO₂ incubator for 24 h.
2. Thaw an aliquot of the lentivirus stock on ice.
3. Add 1 µL of transduction enhance agent into the well with the cells and mix.
4. Add lentivirus to the cells at a multiplicity of infection (MOI) of 5:1 and mix gently. On the next day, repeat this step (24 hours after the first transduction).
5. Monitor the T cell growth every 2-3 days. Add more fresh CAR T-cell medium to maintain the cells at a density of 1 x 10⁶ to 2 x 10⁶ cells/mL.
6. Freeze down the CAR T cells with a standard protocol using a freezing solution (see the Table of Materials).
7. Thaw CAR T cells using a standard method and preculture them in the CAR T-cell medium for about ~2-4 h with interleukin-2, IL-2 (300 units/mL) before applying them to the assay.

3. Detection of CAR expression by flow cytometry

1. Transfer 3 x 10⁵ CAR T cells and nontransduced T cells to two separate 1.5 mL microcentrifuge tubes.
2. Centrifuge the tubes at 300 x g for 2 min and resuspend the cells in 200 µL of fluorescence-activated cell sorting (FACS) buffer containing 1% human serum.
3. Pipet 100 µL of cell solution into two 5 mL polystyrene FACS tubes and keep the tubes on ice for 5 min.
4. Add 1 µL of biotinylated goat anti-mouse F(ab')₂ to one tube of each cell type. Then, add 2 µL of phycoerythrin, a PE-labeled anti-tag antibody (see the Table of Materials) to the other tube of each cell type. Mix well and incubate them on ice for 30 min.
5. Wash the cells with 3 mL of FACS buffer in each tube and centrifuge the tubes at 300 x g for 5 min; discard the supernatants and vortex very briefly or shake the tubes briefly to resuspend the cells in the residual liquid.
6. Add 2 µL of antigen-presenting cells (APC) anti-CD3 and 2 µL of 7-Aminoactinomycin D (7-AAD) antibody solution (see Table of Materials) to each tube. In the tube of cells stained with anti-F(ab')₂ Ab, add 1 µL of PE-labeled streptavidin. Mix briefly and incubate the tubes on ice for 30 more min.
7. Use FACS buffer to wash the cells again as described in step 3.5 and add an additional 200 µL of FACS buffer to each tube.
8. Use flow cytometry to analyze the cells by first gating on the T cells in a forward scatter vs. side scatter plot and then gating on the live cells (7-AAD-negative) in a CD3 vs. 7-AAD plot. The final step is to analyze anti-tag, anti-ScFv, or anti-F(ab')₂ vs. CD3.

4. Real-time cytolysis potency assay (RTCA)

NOTE: Perform the RTCA assay according to the manufacturer's recommended conditions. In brief, first plate the target cells in the wells of the E-Plate, followed by the addition of CAR T cells on the next day. The cytolysis activity of CAR T cells against target cells is monitored in real-time. T cells and mock-transduced T cells (Mock CAR T cells) are used as negative effector cell controls. The following protocol describes an in vitro real-time cytolysis potency assay for adherent tumor cell lines.

1. Add 100 µL of target cell culture medium to each well of the xCELLigence E-Plate, place the plate inside the xCELLigence instrument, and take a background reading. Then, transfer the E-Plate to a tissue culture hood for cell seeding.
2. Use a standard protocol to trypsinize target cancer cells from the culture device. Then transfer the cells to a 15 mL centrifuge tube and add fresh culture medium, up to 15 mL. Pellet the cells by centrifugation for 5 min at 200 x g. Discard the supernatant, add 5 mL of fresh medium, and use a serological pipette to gently resuspend the cell pellet. Count the density of live cells using a hemocytometer and microscope.
3. Adjust the cell density appropriately and then add 100 L of the cell suspension to each well of the E-Plate. The target cell number is typically around 10,000 cells/well for adherent cell lines (BxPC3, Hela-CD19, and SKOV3), or 30,000 cells/well for suspension cells such as Raji (see below for details regarding precoating wells with antibodies in order to tether liquid cancer cells).
4. Equilibrate the E-Plate at room temperature for 30 min to allow the cells to settle evenly on the bottom of the well (this is critical; do not skip this step).
5. Place the E-Plate into the xCELLigence instrument inside the cell culture incubator and start to measure impedance, displayed as Cell Index vs. time, automatically every 15 min.
6. The next day, prepare effector CAR T cells. Make sure to prepare appropriate controls (i.e., Mock CAR T cells, non-transduced control T cells, and/or unrelated CAR T cells) ahead of time to ensure all the cells are ready at the same time. Adjust the effector cells and the control cells to the proper density, and prepare serial dilutions to ensure the desired E:T ratios will be achieved when 100 µL of effector cell suspension is added to each well in step 9.
7. Pause xCELLigence data acquisition and bring the E-Plate from the incubator to a cell culture hood.
8. Remove 100 µL of medium from each well. The amount of residual medium in each well is now 100 µL.
9. Add 100 µL of serially diluted effector CAR T cells or other control cells (i.e. Mock CAR T cells) to achieve the desired E:T ratios.
10. Equilibrate the E-Plate at room temperature for 30 min to allow effector cells to settle, and then place the E-Plate back into the xCELLigence instrument. Resume data acquisition.
11. If so desired, at key time points xCELLigence data acquisition can be paused and the plate removed in order to collect small samples to be analyzed by orthogonal assays (i.e., measuring cytokine production by enzyme-linked immunosorbent assay (ELISA) or flow cytometry).
12. In the EGFR-GITR-CD3 CAR T-cell experiment, measure interferon-gamma (IFNγ) yield with an ELISA kit (follow the instructions of the manufacturer; see the Table of Materials).
    NOTE REGARDING THE USE OF LIQUID CANCERS: For testing nonadherent hematological cancer cells, prior to adding cells the wells of the E-Plate are first coated with an antibody that is specific for an antigen that is expressed on the surface of the cancer cells. For the Raji B cell line, a tethering reagent based on anti-CD40 is used (see liquid tumor killing assay kit in the Table of Materials). Below is the procedure for coating the plate:
13. Dilute the tethering reagent (anti-CD40) with tethering buffer to a concentration of 4 µg/mL.
14. Working inside a tissue culture hood, add 50 µL of the diluted tethering reagent to each well of the E-Plate. Leave the E-Plate at room temperature or in a 37 °C incubator for 3 h.
15. Remove the tethering reagent and wash the E-Plate at least 2x with wash buffer. At this point, the E-Plate is ready for seeding the Raji target cells (30,000 cells/well).
16. Continue with step 4.1 (above) to perform the rest of the procedure.