Hi,

I have been isolating neutrophils for N-Formylmethionyl-leucyl-phenylalanine (FMLP) activation for a couple of months without success.

In my protocol,

• I dilute heparinized whole blood with PBS (without Ca2+ and Mg2+) in a 1:1 ratio.
• I then layer the blood over Ficoll Pague (1.077) and centrifuge at 600 rcf for 15 min.
• The plasma, peripheral blood mononuclear cells (PBMC) and Ficoll Pague are discarded leaving the red blood cells (RBCs) with granulocytes.
• The RBCs are resuspended in PBS (without Ca2+ and Mg2+) and immediately mixed with 3% Dextran solution 4 oC.
• Sedimentation is allowed for 30 min at 4 oC after which the neutrophil rich supernatant is centrifuged at 600 rcf for 5 min.
• The remaining RBCs are lysed with 155 mM NH4Cl, 10 mM KHCO3, 0.1 mM EDTA for 20 seconds.
• Lysis is stopped with PBS (without Ca2+ and Mg2+), centrifuged at 400 rcf for 10 min at 4oC, and washed with PBS (without Ca2+ and Mg2+) at 4oC.
• The final neutrophil pellet is resuspended in RPMI 1640 and kept on ice.

For flow cytometry,

• I treat 250,000 cells with FMLP 1 uM and 0.2% DMSO for 30 min at 37 oC, 5% CO2. Staining is done with CD11b and isotype control for 15 min at room temperature (RT) in the dark, after which the cells are washed twice with PBS to remove any unbound antibodies, and then fixed with 1% paraformaldehyde (PFA) for 30 mins before measurement.

In my results,

• I observe no differences in the fluorescence intensity between FMLP and DMSO. This experiment has been repeated a few more times without success.
• Interestingly, I see differences in the fluorescence intensity between the two groups when I use whole blood instead of isolated neutrophils, but my interest is in the latter.
• What could be the reason for these failures? Is my isolation or staining protocol wrong?