Abstract
A method for Ca2+ flux measurement on isolated human peripheral B cells that uses flow cytometry is described. B cells were isolated by anti-CD19 magnetic bead sorting, and Ca2+ flux was measured with the fluo-3 reagent on a standard single-laser flow cytometer. The response of B-cell stimulation by anti-immunoglobulin B (anti-IgM), anti-IgD, protein A, concanavalin A, and ionomycin was determined. Percentage of responder B cells, the level of Ca2+, and the time of peak stimulation were measured. Bound anti-CD19 monoclonal antibody coupled with small paramagnetic particles did not affect Ca2+ flux. All the isolated B cells responded maximally at 10s with stimulation by 8 microg of ionomycin. The average isolated preparation contains 70% IgM+ and 85% IgD+ cells, all of which showed peak stimulation with 10 microg of anti-IgM and anti-IgD per ml, respectively, at 30s. Only at high concentrations of 80 microg/ml, concanavalin A produced a slower response, peaking at 90 s after stimulation. Stimulation with 20 microg of protein A per ml resulted in Ca2+ flux in only 40 to 60% of cells that had a rapid response and maximal stimulation resembling the pattern of activation of ionomycin. B cells from three patients with mixed cryoglobulinemia with high concentrations of monoclonal rheumatoid factors showed stimulation with aggregated IgG, whereas those from healthy control subjects did not, demonstrating the applicability of the methodology to detection of specific antigen stimulation of B cells. This methodology may be useful in testing the functional capacity of B cells in a variety of diseases. The methodology may also prove useful in studying antigen-specific B-cell responses when they involve a significant percentage of B cells.
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Selected References
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