Step 1:  Count cells so that there are 5 x 10^5 – 5 x 10^6   cells per sample ( i.e. per well or per tube ).  Transfer each sample to a round-bottom 96-well plate or 12x75mm collection tube.

Step 2:  Centrifuge in the cold at 1200 rpms for 3 – 5 minutes; remove supernatants, vortex, and place on ice.Step 3:  Add 25 microliters/well or 100 microliters/tube of appropriate antibody (primary antibody, direct fluorochrome-conjugated antibody, or fluorescent isotype [control] diluted to the correct concentration as determined by optimization experiment for each) in staining buffer, or staining buffer alone (unstained control), to each well.

Step 4:  Gently vortex and incubate on ice in the dark for 30 minutes.

Step 5:  Wash plate/tubes 2 – 3 times with staining buffer:  fill each well with 0.2 ml of staining buffer (1 ml for tubes), centrifuge for 5 minutes in the cold, remove supernatant, VORTEX, repeat.

Step 6:  Repeat steps 3-5 with secondary antibody, if needed.

Step 7:  Resuspend in 0.2 ml of staining buffer, transfer to 5 ml tubes (12 x 75 mm) – if staining in plates, and add 0.2-0.3 ml of staining buffer or paraformaldehyde. Vortex and place on ice in the dark until analyzing in the UAMS Flow Cytometry Core Facility.

Note:  When fixing samples with paraformaldehyde as the last step, it is very important to VORTEX cells on high to break up the pellet of cells before adding the paraformaldehyde.  Once samples have been fixed with paraformaldehyde they may be stored in the refrigerator overnight (or up to one week) before being analyzed on the flow cytometer.