Multiparametric flow cytometry and CATALYST analysis

1. Disassociation Buffer (20mL):

*The use/amount of Dispase II is lot dependent.

3. Place tubes on ice and bring with you to the procedure room for animal take down and sample collection.

4. Sack mice one by one using CO₂ and retrieve sample, place each sample in their respective conical tube on ice.

5. Once all tissue are collected place the tubes with the tissue and disassociation buffer in the shaking incubator at 200rpm and 37℃ for an hour.

Sample Processing:

1. Following the 1h incubation transfer the samples into a BSL-2 hood to continue sample processing.
2. Transfer the tissue and disassociation volume buffer on-top of the cell-strainer FACS tubes. (ie. small amount of volume, tissue, volume repeat until fully transferred).
3. Using a 3ml syringe plunger gently muddle the tissue remaining on the strainer. DO NOT BREAK THE FILTER.
   1. You may use the same syringe plunger for the same conditions.
   2. Tapping the tube will help pass liquid through the filter.
4. Spin the tubes using the programmed flow wash setting on the TC Centrifuge (350g for 5 minutes), this program will be used throughout the flow staining protocol for each centrifuge run. Following the 5 minutes decant the tubes supernatant into the sink and then proceed to do one wash with 5mL of PBS and spin down.
   1. This helps remove any residual fat cells.
5. 10x RBC has to be diluted to 1x w/ the ultra pure distilled water (100µL 10X RBC to 900 µL UltraPure Distilled Water).
6. 2mL of the 1x diluted RBC buffer is added to each tube and reacted on ice for 5 minutes, do not resuspend.
7. Add 2mL of FACS to the RBS to stop the reaction and then spin down and decant.
8. Add 2mL of FACS for a wash and spin down and decant.
9. Add 2mL of PBS to wash and remove residual FBS and pass through new strainer FACS tubes and then spin down and decant.
   1. The PBS wash helps remove residual FBS as FBS will inactivate zombie violet dye.
   2. This second pass through a cell strainer helps remove any cell aggregates before beginning the dying process.

Dying Process (Lymphoid and Myeloid Breakdown):

1. Make tubes for single stain controls and unstained controls for both panels (this includes a single stain tube for Zombie Violet dye).
   1. If you do not have a large pellet, you can spin down the sample tubes you transferred through another strainer to collect the residual fluid for the controls.
2. Transfer a small volume (~20 ul) of cells to each control tube made (some stains such as CD206, PDL1, CD25, PD-1, CD223, CD69, CD103, CCR7, Granzyme B, Perforin, and FOXP3 should be taken from conditions that have PDL1 protein presentation).
3. Make a 1: 100 dilution of zombie violet for all sample tubes and the two-zombie violet single stains. (Ex. 25μL of Zombie Violet into 2475µL of PBS for 25 tubes)
4. Add 100uL of the Zombie Violet 1:100 dilution to each tube, vortex and place them covered from light in the 4℃ for a 15-minute incubation.
   1. Single stains- apart from the zombie violet single stain- will not receive the zombie violet dye
5. During the incubation make the antibody cocktails for the two respective panels. Each sample will get all the dyes (apart from the lymphoid panel which will receive every dye but the PE FoxP3 until the permeabilization step).
   1. Make antibody cocktails with a two-tube buffer (ie. 22 tubes for 20 samples)
   2. Each tube will receive 1µL of each antibody (unless stated otherwise in titration) into the ~100µL of volume that is left in the tubes following decanting. This total will match the number of antibody dyes you are using within the panel. (Ex. The total volume added will be 12µL for the Myeloid Panel Dye Cocktail and 9µL for the Lymphoid Panel Dye Cocktail.)
   3. For the lymphoid panel two ab cocktails should be made. FoxP3, Perforin, and GranzymeB (intracellular dyes) and all other dyes (extracellular dyes)
   4. Single Stains and FMOs will get 1 ul of the respective dyes rather than the antibody cocktail.
6. Following the 15-minute incubation for the zombie violet dye add 1mL of FACS for a wash, spin down, and decant.
7. Transfer half of your samples (ie. if 80µL of volume 40µL will be transferred) to create your tubes for the Myeloid Panel. This will separate the tubes into the two respective panels as they will be treated separately throughout the rest of the protocol.
8. Add FACS to each tube to bring up to 100µL. (ie. 60µL of FACS is 40µL of volume was moved).
9. True Stain or True Stain Monocyte + True Stain Treatment:
   1. For the Lymphoid Panel: 1µL of true stain should be added to each tube, vortexed, and left in the 4℃ covered to incubate for 15 minutes.
   2. For the Myeloid Panel: Add 5µL of True Stain Monocyte to each tube and then add 1µL of True Stain and vortexed. Leave to sit and incubate at 4℃ covered for 15 minutes.
   3. NOTE FOR BOTH: At this step also transfer volume of the samples to create FMOS (take small volumes ~20 ul from conditions with PDL1 present).
10. Following the incubations add stain cocktail to all the samples (respective to the panels) and add the stains to the Fluorescence Minus One (FMOs) and Single Strain controls. Vortex each tube, cover the racks and leave in the 4℃ for a 30-minute incubation period.
    1. FMOs are dye specific which will include every dye except the one of interest, Ex. a FoxP3 FMO will have every dye except the one for FoxP3. These are used to help determine your gates for your populations of interest.
11. During the incubation make the solutions required for nuclear staining from the nuclear stain kit: Note: You should be calculating out how much you need for the number of tubes you have and then making the solutions based on the dilutions stated below.
    1. Make a 1:10 dilution of the 10X Permeabilization Buffer with Ultrapure Distilled water to obtain a 1X solution to add to the tubes. Each tube will receive 2mLs.
       1. (ie. for 37mLs 3.7mL of 10X Permeabilization Buffer will be added to 33.3mL of UltraPure Distilled water).
    2. Make a 1:4 dilution of the 4X Fixation Buffer provided in the kit.
       1. (ie. for 10mL add 2500µL of the 4X Fixation Concentrate to 7500µL of the Fixation dilutant provided in the kit).
12. After the incubation period all tubes are washed with 2mL of FACS and spun down and decanted.
13. Fixation:
    1. For the Lymphoid Panel: Add 200µL of the 1X Lymphoid Fixation Buffer you made from the nuclear staining kit and let sit for 30 minutes as 4℃ covered.
    2. For the Myeloid Panel: Add 500uL of Fixation Buffer (IT IS NOT THE SAME ONE FROM THE NUCLEAR STAIN KIT) and let it incubate for 20 minutes at room temperature.
14. Once the Myeloid Incubation period is up spin down the tubes, decant,  add 2ml of FACs for a wash, spin down and decant. Once the FACs wash is complete these samples can be stored at 4℃ stained to be read day of or the following day.
15. Once the Lymphoid Panel incubation is up add 1mL of the 1X Permeabilization buffer to each tube and spin down and decant.
16. Following the permeabilization buffers the samples and tubes that receive intracellular dyes will receive the intracellular ab cocktail to each sample. These will be then vortexed and left to incubate at 4℃ for 20 minutes protected from light.
    1. These tubes include ALL ANIMAL SAMPLES, FoxP3, Perforin, and Granzyme B Single Stains, and all FMOS with the respective dyes
17. After the 20 minute staining incubation add 1mL of the 1X Permeabilization buffer to all tubes (including the ones without FoxP3 dye). Spin down and decant.
18. Once decanted add 1mL of the FACs buffer for the final wash and then spin down and decant.
    1. These can be left after this step at 4C to be read another day.
19. Sample analysis was performed using the CATALYST analysis tool. A github from the software author with detail explanation is available here https://github.com/HelenaLC/CATALYST.