Assessing T-cell Receptor-Induced Calcium Influx Using a Calcium Indicator Dye

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Preparation of immune cells from the mouse spleen

NOTE: Euthanize naïve mice with carbon dioxide (CO₂) euthanasia. C57BL/6 mice purchased from Jackson Laboratories and bred in-house are used for experiments at 6-12 weeks of age. Both male and female mice are utilized for experiments.

1. Disinfect the mouse skin with 70% ethanol in order to reduce the possibility of external contaminants from getting into the sample.
2. Dissect the mouse spleen using dissection tools, surgical scissors, and forceps. Harvest the spleen and place the detached spleen into a 50 mL conical tube with ~5 mL of complete Roswell Park Memorial Institute media (cRPMI) on ice. 
   NOTE: Complete RPMI is comprised of 10% fetal bovine serum (FBS), 2 mM L-glutamine, and 1% penicillin/streptomycin.
   1. To harvest the mouse spleen, make a 5 cm incision into the fur and skin along the left side of the mouse halfway between the front and back legs with scissors. Open the body cavity ~5 cm and remove the spleen using forceps. The spleen is the color of a kidney bean and is longer and flatter than the neighboring kidney.
3. Pour the contents of step 1.2 onto a sterile 70 µm filter attached to a new 50 mL conical tube. Mechanically separate the spleen into a single-cell suspension by pushing the spleen through the filter with a 5 mL syringe plunger until no spleen pulp remains on the filter surface.
4. Pellet the splenocytes utilizing a swing-out rotor countertop centrifuge at 500 x g for 5 min at 4 ˚C.
5. Carefully decant the supernatant. The pelleted splenocytes remain at the bottom of the 50 mL conical tube.
6. To remove red blood cells, re-suspend the pelleted splenocytes in 1 mL of ammonium–chloride–potassium (ACK) lysis buffer for 3 min according to the manufacturer's instructions. Mix well.
7. Quench the ACK lysis buffer with 15 mL of cRPMI.
8. Pellet the lymphocytes as directed in step 1.4.
9. Re-suspend the cell suspension in 5 mL of cRPMI in a 50 mL conical tube. Place the samples on ice.
   1. To count the cells, transfer 10 µL of the cell suspension to a 0.6 mL microcentrifuge tube and dilute at a 1:1 ratio with Trypan blue solution. Then, transfer to a hemocytometer or automated cell counting system.
10. After counting, depending on the cell concentration in cell suspension as in step 1.9, pellet the cells in a tabletop centrifuge at 500 x g at 4 ˚C for 5 min to prepare for the addition of cRMPI media containing Indo-1 AM ester dye.
11. Calculate the volume for re-suspension of the cells to achieve 10-12 x 10⁶ cells/mL. This is a 2x concentration of the total cell count required.
12. Re-suspend the cells in the volume of cRPMI calculated in step 1.11. Place the cells at 37 ˚C with 5% CO₂ in either a 50 mL conical tube with the lid vented or a tissue culture plate.

2. Indo-1 ratiometric dye and fluorescent antibody labeling

1. As per the manufacturer's instructions, add 50 µL of dimethyl sulfoxide (DMSO) into a vial containing 50 µg of Indo-1 AM. The concentration of the stock solution is 1 µg/µL.
   NOTE: DMSO is provided in micro-vials with the kit to prevent any oxidation of DMSO.
2. Dilute the stock aliquot (1 µg/µL) into the appropriate experimental volume (established in 1.11) of cRPMI at a concentration of 6 µg/mL, a 2x concentration.         Do not store Indo-1 in an aqueous solution.       
   NOTE: Other buffers with added calcium can be used depending on the cell needs.
3. Set complete media with Indo-1 aside in a water bath at 37 ˚C.    
   NOTE: Use the minimum concentration of Indo-1 AM ester necessary to obtain an adequate signal. This needs to be titrated for varying cell types. Typically, a concentration between 3-5 µM is sufficient. For primary T-cells, loading cells with Indo-1 at a concentration >5 µg/mL increases mean fluorescent intensity (MFI) above a log of 6 on spectral flow cytometers. If this occurs, decrease ultraviolet (UV) laser intensity and titrate the Indo-1 to a lower concentration.
4. Dilute previously prepared cell suspension (in step 1.12), 1:1 in cRPMI, including the 2x Indo-1 media made in step 2.2. Ensure that the cells are at a concentration between 5-6 x 10⁶/mL.
   NOTE: Do not dye load unstained single stain control or surface marker single stain cell controls with Indo-1. Adding Indo-1 to single stain antibody controls will create a multi-color sample due to the Indo-1 added to single stained cells. Include indo-1 (calcium-free) ethylene glycol tetraacetic acid (EGTA) added control to the sample plate created in step 2.6.
5. Add 1 mL of cells/well/experimental condition into a 12-well tissue culture plate.
6. Create a single stained control for Indo-1 (calcium-free) sample using previously Indo-1 dye-loaded cells and add EGTA to a final concentration of 2 mM. EGTA will chelate calcium in the cell suspension, giving a cleaner control sample.
   1. Create an Indo-1 positive control (Indo-1 calcium-bound) by adding 50 µM of ionomycin to the dye loaded cell suspension at the flow cytometer. Ionomycin is a membrane permeable calcium ionophore that binds calcium ions and facilitates the transfer of calcium ions into cells.
7. Create a well for biological negative control with no fluorophores or Indo-1 dye.
8. Create a well for single stain controls for any additional cell populations of interest (antibodies user defined) using antibody conjugated fluorophores in a flow cytometry panel design. These will not include the Indo-1 ratiometric dye.
9. Add the Fc receptor blocking antibody (~2 µg/1 x 10⁶ cells), as per the manufacturer's instructions, in advance of adding additional fluorochrome-conjugated antibodies for surface staining.
10. Incubate the plate for 45 min at 37 ˚C with 5% CO_2.
11. Resuspend the cells in a 12-well plate by gently tapping the plate every 15 min.
12. Mix and remove the entire volume of cells suspended in the media from the 12-well plate. Then add the cell suspension to 1.7 mL microcentrifuge tubes.
13. Pellet the cells in a microcentrifuge at 500 x g for 5 min at room temperature. Decant the supernatant and wash the cells by adding 1 mL of pre-warmed cRPMI. Pellet again and decant the supernatant.       
    NOTE: Washing the cells removes any remaining Fc blocking antibody.
14. Resuspend the cells in 1 mL of cRPMI in 1.7 mL microcentrifuge tubes and incubate each tube at 37 ˚C for an additional 30 min at 5% CO₂, leaving the top of the tube slightly open to allow for gas exchange. This allows complete de-esterification of the intracellular AM esters.
15. Transfer the cells back into a 12-well tissue culture plate, if needed. Additional user defined titrated fluorochrome-conjugated antibodies can be added at a resting phase.
    NOTE: Markers used in the methods panel include: Ghost 540, CD4, CD8, TCRβ, TCRγδ, CD25, and CD1d/α-galcer tetramer. Markers are chosen to analyze T-cell subsets.
    1. Create a master mix of all the user-defined fluorochrome-conjugated antibodies according to the cell types of interest at a previously titrated volume into a 1.7 mL microcentrifuge tube.
    2. Add a calculated master mix for 1 mL of the cell volume, dependent on titrated antibodies to resting cells.   
       NOTE: The advantage of staining at step 2.15 instead of the later step 2.18 is that staining at step 2.15 will decrease the overall incubation time for the experiment. However, staining in a total volume of 1 mL at step 2.15 increases antibody usage.
16. After rest, pellet the cells at 500 x g for 5 min at room temperature.
    NOTE: Cells in a 12-well tissue culture plate can be pelleted in the plate with a plate centrifuge accessory. Otherwise, pellet the cells in a 1.7 mL microcentrifuge tube.
17. Wash the cells by resuspending the pellet in 1 mL of 4 ˚C cold cRPMI and pellet the cells again as in step 2.16. Place the cells on ice.   
    NOTE: If surface staining cells separately, post de-esterification at step 2.18, less antibody volume is needed. Surface staining at this step can be performed after the rest phase in cold flow buffer (1x Dulbecco's phosphate-buffered saline (DPBS) with added 1% FBS), using user-defined antibodies, on ice for 30 min. Wash the cells after the stain in cRPMI as in step 2.17.
    1. Add viability stain Ghost540 diluted 1:7,500 in DPBS to samples according to the manufacturer's instructions. Heat kills the cells at 55 ˚C on a warming block for single stained live/dead control.         
       NOTE: Viability functional dye staining to eliminate dead cells in flow cytometric analysis is performed without FBS. FBS can interact with amine dyes as per the manufacturer's instructions.
18. Re-suspend individual samples in 500 µL of cRPMI (phenol red-free) using a 5 mL polystyrene flow tube with a cap.        
    NOTE: Cells can be left at 4 ˚C for up to an hour.
19. Warm every 5 mL tube containing cells, individually, to 37 ˚C using a bead bath for 7 min prior to the analysis on the flow cytometer, keeping time consistent between the samples. Use a timer.

3. Bead bath tube: maintaining temperature during calcium flux analysis

1. Add bath beads to a small water bath with no water added; warm up to 37 ˚C.
2. Carefully cut a 50 mL conical tube in half, at the 25 mL mark, with a razor blade; discard the top of the tube.
3. Fill the halved tube ¾ of the way with bath beads.
4. Place the tube created in step 3.2 into the bead bath created in step 3.1. Bring the tube and beads to 37 ˚C.
5. Plug the bead bath into an electrical outlet next to the flow cytometer in order to maintain the temperature in preparation for sample analysis.
6. Add a 50 mL conical Styrofoam rack cut to hold one tube to position the tube in place while running on the flow cytometer. This will eliminate the need to manually hold the tube for the duration of the curve analysis.