Flow Cytometry Protocols

• Flow cytometry works with cells at a concentration of 10⁵ – 10⁷ cells/ml. More concentrated means that the cytometer may clog
• Cells that are already in solution are the easiest to run, but adherent cell culture and solid tissue can be run once they are dispersed into a single cell solution by enzymatic and/or mechanical means.
• Enzymes should be carefully optimized, depending on pH, presence of divalent cations, tissue type etc.
• For intracellular antigens, cells must be permeabilized and fixed to allow access for the antibody, to maintain the cell’s integrity and preserve labile components

Cultured Cells in Suspension

1. Transfer cultured cells to conical centrifuge tubes.
2. Centrifuge cells for 5 min at room temperature at 300-400 g.
3. Discard supernatant and resuspend cells in 10 ml PBS containing 1% BSA.
4. Repeat steps 2 and 3, this time resuspending in cold (4°C) PBS-BSA to a concentration of ~1 x 10⁶ cells/ml.

Peripheral Blood Mononuclear Cells

Note that whole blood can be used directly in flow cytometry, though red blood cells are typically lysed or removed first. The procedure below isolates PBMCs from blood.

1. Equilibrate separation media, such as Ficoll, to room temperature.
2. Dilute blood sample in an equal volume of PBS containing 1% BSA.
3. Transfer a volume of separation media equal to the volume of diluted blood into a 15 ml conical tube.
4. Carefully overlay the blood onto the separation media in the tube.
5. Centrifuge at 300–400 g for 30 min at 20°C in centrifuge with no brake on.
6. Harvest the PBMC layer using a pipette, typically between plasma and the separation media. Transfer harvested cells to a 15 ml conical tube.
7. Add PBS-BSA to the PBMCs to a volume of 10 ml. Centrifuge at 300–400 g for 5 min at room temperature.
8. Discard the supernatant, and resuspend the PBMCs in cold (4°C) PBS-BSA to a concentration of ~1 x 10⁶ cells/ml.

Cultured Adherent Cells

1. Remove cell culture media, rinse in PBS, and detach the cells from their tissue culture flask or plate by cell scraper, 0.25% trypsin, or Accutase.
1. Quench enzymes by adding media containing serum.
2. Gently triturate to disperse cell clumps.
2. Transfer to a conical tube and centrifuge at 300–400 g for 5 min at room temperature.
3. Resuspend in PBS-BSA and centrifuge again at 300–400 g for 5 min at room temperature.
4. Resuspend in a small volume of cold PBS-BSA.
5. Count the cells with a hemocytometer, and adjust the concentration to ~1 x 10⁶ cells/ml with more PBS-BSA.

Tissue

1. Dissect and harvest the desired tissue
2. For lymphoid tissue (spleen, lymph nodes, thymus):
1. Transfer tissue into a tissue culture dish containing 10 ml PBS-BSA.
2. Disperse the tissue into a single cell suspension by mashing with the plunger of a 3 ml syringe, or with the frosted ends of two microscope slides.
3. For non-lymphoid tissue:
1. Mince into 2-4 mm pieces using scissors or scalpel.
2. Dissociate the tissue into a single cell suspension. This will vary depending on the tissue, but will usually require incubating the tissue with enzymes (e.g. collagenase, dispase, DNase etc.) at an optimal temperature (e.g. 37°C) for the appropriate amount of time (often 10-30 minutes). Optimizing this step is important to ensure good recovery and viability of samples.
4. Pass cells through a nylon mesh cell strainer into a 15 ml conical tube to remove clumps or debris.
5. Centrifuge at 300-400 x g for 5 minutes at 4°C.
6. Discard the supernatant and resuspend in 10 ml PBS-BSA.
7. Centrifuge at 300-400 x g for 5 minutes at 4°C.
8. Repeat steps 6 and 7. Note that this amount of washing may not be necessary for all tissue types.
9. Resuspend in a small volume of cold PBS-BSA.
10. Count the cells with a hemocytometer, and adjust the concentration to ~1 x 10⁶ cells/ml with more PBS-BSA.

Direct staining refers to using a primary antibody that has been directly conjugated to a fluorophore, while indirect staining requires a conjugated secondary antibody to detect an unlabeled primary antibody.

This protocol is applicable to cell suspensions as prepared above, or whole blood. In the case of whole blood, an anticoagulant such as heparin, EDTA or sodium citrate is required.

1. Adjust cell suspension to a concentration of ~1 x 10⁶ cells/ml with PBS-BSA. Whole blood may need to be diluted if the cell count is high, such as in leukemia samples. An anticoagulant will also be necessary for blood.
2. Aliquot 100 μl of the sample into as many test tubes as required.
1. Be sure to aliquot enough for all of the required Flow Cytometry Controls.
3. Add primary antibody at an optimized or recommended dilution. Mix well and incubate at 4°C for at least 30 minutes. Avoid direct light if using fluorescent conjugates.
4. For blood samples: add 2 ml 1X red blood cell lysis buffer and mix well. Incubate until the sample is clear (10-15 minutes). Centrifuge at 300–400 g for 5 min at room temperature and discard the supernatant.
5. Wash cells by adding 2 ml cold PBS-BSA to cells and centrifuge at 300–400 g for 5 min at 4°C. Discard the supernatant.
6. For indirect staining: Dilute secondary antibody at the recommended dilution in PBS-BSA and add to cells. Mix and incubate at 4°C for at least 30 min. Avoid direct light.
7. Resuspend in 200 μl cold (4°C) PBS and run samples through flow cytometer as soon as possible. Avoid light and keep cells on ice.
1. If flow cytometry analysis will not occur for more than 1 hour or biohazardous agents need to be inactivated, fixation may be necessary.
2. Fixation can be achieved by treating with 100 μl 0.1-1% paraformaldehyde for 10-15 minutes on ice, 1 ml ice cold acetone on ice, or 1 ml methanol at -20°C for 5-10 minutes. Centrifuge and wash twice in PBS-BSA after fixation.
3. Cells should not be fixed if they must remain viable.

Staining for intracellular antigens requires additional fixation and permeabilization steps so that antibodies can access their targets while preserving cell morphology. When detecting cytokines, pre-treatment with monensin (1-3 µM) or brefeldin A (5 µg/ml) for 4-8 hours is usually necessary to inhibit secretion. Stimulation of cytokine production for positive controls, optimization or other experimental aims, may be achieved using phorbol myristate acetate (PMA; 10 ng/ml) and ionomycin (2 μM) alongside monensin/brefeldin A treatment.

This protocol is applicable to cell suspensions as prepared above, or whole blood. In the case of whole blood, an anticoagulant such as heparin, EDTA or sodium citrate is required.

1. Adjust cell suspension to a concentration of ~1 x 10⁶ cells/ml with PBS-BSA. Whole blood may need to be diluted if the cell count is high, such as in leukemia samples. An anticoagulant will also be necessary for blood.
2. Aliquot 100 μl of the sample into as many test tubes as required.
1. Be sure to aliquot enough for all of the required Flow Cytometry Controls.
3. If also staining for cell surface markers: Staining of cell surface markers must be performed before fixation and permeabilization. Add fluorescently conjugated primary antibody at an optimized or recommended dilution. Mix well and incubate at 4°C for at least 30 minutes. Avoid direct light.
4. Wash cells in 2 ml PBS-BSA at 4°C. Centrifuge and discard the supernatant.
5. Fix the cells: resuspend cells in 100 μl fixative per 1 x 106 cells for 10-20 minutes. Mix well to avoid cell clumping. This step will require optimization and depends on the antigen, which may be masked by certain fixatives. Some suggested fixatives and guidelines for treatment conditions are below:
1. Paraformaldehyde (PFA), 1-4%, 15-20 minutes on ice
2. Methanol, 90%, 10 minutes at -20°C
3. Acetone, 100%, 10-15 minutes on ice
6. Wash the cells twice: centrifuge at 300-400 g for 5 minutes at 4°C, then discard the supernatant and resuspend in excess PBS-BSA.
7. Permeabilize the cells: centrifuge at 300-400 g for 5 minutes at 4°C, then discard the supernatant and resuspend in detergent of choice for 10-15 minutes at room temperature.
1. This step is not required if using acetone or methanol for fixation, which also permeabilize cells.
2. Permeabilization affects light scatter during cytometry, which may affect gating settings.
3. The optimal detergent will depend on the specific antigen and its localization in the cell:
1. Strong detergents, including Triton X-100 and NP-40, penetrate organelle membranes including the nuclear envelope, allowing nuclear staining. Use at 0.1-1% in PBS.
2. Weaker detergents, including Tween-20, saponin and digitonin, are mainly suitable for antigens in the cytoplasm rather than in organelles. Use at 0.1-0.5% in PBS.
3. Methanol can also be used as a permeabilization agent following PFA fixation. Use at 90% on ice. Methanol denatures protein-based fluorophores such as PE, so do not use methanol if these fluorophores have already been used (e.g. to stain cell surface markers).
8. Wash the cells twice: centrifuge at 300-400 g for 5 minutes at 4°C, then discard the supernatant and resuspend in excess PBS-BSA.
9. Add primary antibody at an optimized or recommended dilution. Mix well and incubate at 4°C for at least 30 minutes. Avoid direct light if using fluorescent conjugates.
10. Wash the cells twice: centrifuge at 300-400 g for 5 minutes at 4°C, then discard the supernatant and resuspend in excess PBS-BSA.
11. For indirect staining: Dilute secondary antibody at the recommended dilution in PBS-BSA and add to cells. Mix and incubate at 4°C for at least 30 min. Avoid direct light. Wash the cells twice after incubation as above.
12. Resuspend in 200 μl cold (4°C) PBS and run samples through flow cytometer as soon as possible. Avoid light and keep cells on ice.

The cell cycle status of an experimental sample can be determined by measuring the quantity of DNA present in each cell. Propidium iodide (PI) and other dyes such as DAPI, Hoechst and 7-AAD bind to DNA in proportion to the DNA quantity, and so can distinguish between different phases of the cell cycle by the amount of DNA present: G0/G1 phase (2n), S phase (2n~4n), and G2/M phase (4n).

To distinguish G0 from G1, staining for Ki-67 may be performed, which is a marker of proliferating cells and rarely detected in G0 phase. Experiments involving incorporation of the uridine analogs BrdU and EdU can provide more information on cell cycle status kinetics.

Cells must be fixed and permeabilized to allow the dye to access the DNA in the nucleus. Alcohol such as ethanol is commonly used for this purpose, which fixes and permeabilizes cells simultaneously, but alcohol may not be compatible with other stains or fluorescent proteins. If co-staining with incompatible markers is required, PFA and a permeabilization agent such as Triton X-100 can be used instead.

1. Prepare cell suspension as described in Sample Preparation.
2. Centrifuge cells at 300-400 g for 5 minutes at 4°C, then discard the supernatant.
3. Add cold (4°C) 70% ethanol dropwise to the cell pellet while gently vortexing. Fix cells for 30 minutes on ice.
1. a. 70% ethanol should not be made with PBS because this causes protein precipitation during fixation. Make with distilled water instead.
4. Centrifuge at 500 g for 10 min. Discard supernatant.
5. Wash the cells twice: centrifuge at 300-400 g for 5 minutes at 4°C, then discard the supernatant and resuspend in excess PBS.
6. Pellet the cells again and discard the supernatant. Add 50 μl PBS containing 100 μg/ml RNAse A to prevent staining RNA.
7. Add 400 μl PI solution (50 μg/ml in PBS) per 1x106 cells directly to cells in the RNase A solution. Mix well and incubate for 5-10 minutes at room temperature. Some cell types may require a longer incubation time.
8. Analyze on flow cytometer.
1. Identify cells using forward and side scatter.
2. Exclude doublets use pulse shape analysis.
3. Analyze cell cycle populations using a one-parameter histogram of PI signal.