I am trying to look at Calcium Fluxes in primary mouse T cells after activation (with peptide on DCs, CD3/CD28, etc). I am looking to do this study on a microscope and have been using Fluo-4 AM (Invitrogen). My method (see below) successfully works for a B cell line. However, when I stain primary mouse T cells, my cells look very green even before activation and hence activation (even positive control with ionomycin) yields a very small signal that I cannot even detect with any certainty. It seems like the cells take up so much dye initially, that there is hardly any further increase in signal after activation. Maybe they also compartmentalize the dye (?). In any case, I have also tried to use only 10 minutes of incubation time (at room temperature, rather in the incubator) with 0.25 MicroM Fluo-4 - this is four times lower than the minimum time and concentrations recommended by invitrogen. But even with this method I cannot detect any change in fluorescence.
Has anyone in this forum done Calcium Flux measurements with primary mouse T cells before and would mind sending me their protocol (ideally for microscopic studies; FACs prep should be similar as well, though)? Else, do you have any idea of what might be going wrong with my primary T cell staining or do you have any recommendations?
I would appreciate any help greatly - that would be truly wonderful! Thanks so much and many kind regards xxx
dissolve Fluo-4AM (50microgram) in 25 microliter anhydrous DMSO
stain 2Mio primary T cells/ml in HBSS (no Phenol Red), 1% Serum, 3mM Probenecid and 0.5 microM Fluo-4 AM for 30 minutes at room temperature
wash the cells twice with HBSS (includes 1mM Calcium) and 1%Serum. Incubate for another 20 minutes in the incubator.
Activate with Ionomycin (2-20microM) or CD3/CD28
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Calcium Flux Primary Mouse T Cells (Fluo-4)
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