Isolation and Activation of CD4+ T and B Cells

Primary human CD4⁺ T and B cells were isolated from the blood of healthy human adults, as described above. Cells were seeded at ~1 × 10⁶ cells/mL in RPMI-G on a TC-treated 96-well plate (180 μL in each well). CD4⁺ T cells were stimulated with immobilized anti-human CD3ε (0.5 μg; BD Biosciences) and PMA (10 ng/mL; Sigma), as described^{46 (link),47 (link)}. B cells were stimulated with human IL-4 (20 ng/mL; Sigma) and anti-human CD40 monoclonal antibody (5 μg/mL; Enzo, clone mAb 89), as described^{48 (link)}. Immediately following stimulation, cells were treated with either PBS or R-P4 (20 μM) in technical triplicate. As controls, a group of CD4⁺ T and B cells received no stimulus and no treatment (designated as “PBS (unstimulated)”). After incubating for 48 h at 37 °C, cells were treated with human Fc block (1:200; BD Biosciences), stained with anti-CD69-PE/Cy7 (10 μL/test; BD Biosciences, clone FN50), washed, and resuspended in DAPI (0.5 μM; Thermo). Cells were run on an LSR II flow cytometer (BD Biosciences). Unstained cells, single-stained DAPI+ cells, and single-stained PE/Cy7+ beads were used as compensation controls. Data were analyzed using the FlowJo software, and gates for DAPI− and CD69+ events were determined using fluorescence minus one or unstained controls. For the gating strategy used to calculate percent CD69⁺ cells of single cells, see Supplementary Fig. 5.