Up-regulation of GITR and CTLA-4 following IL-2 administration. (A) Purified CD4 T cells or PBMCs were stained with allophycocyanin-conjugated anti-CD4, PE-conjugated anti-CD25, and FITC-conjugated anti-CD69, anti-GITR, or (C) anti-CD27 antibodies. Dot plots were gated on CD4⁺ T cells. (B) Purified CD4 T cells or PBMCs were stained initially with either FITC-conjugated anti-CD4 or anti-CD8 antibodies with allophycocyanin-conjugated anti-CD25 antibody, followed by intracellular staining with PE-conjugated anti-CTLA-4 antibody. The dot plots were gated on CD4⁺ or CD8⁺ T cells, respectively. The quadrants for CD69, GITR, CD27, and CTLA-4 were based on the isotype control antibodies as well as T cells that lack expression of these markers. This result is representative of 6 patients and 3 healthy donors.

IL-2 increased the frequency of CD4⁺CD25⁺Foxp3⁺ T cells in peripheral blood. (A) RNA was extracted from highly purified CD4 T cells (> 97%-99%) isolated from PBMCs and collected through leukapheresis of 8 patients pretreatment (PRE), during rebound phase (REB), and 2 to 3 weeks after treatment (POST). Foxp3 levels were quantified using real-time RT-PCR and normalized for endogenous β-actin. The P value was calculated using paired t test and adjusted for multiple comparisons. (B) To determine Foxp3 expression per cell, cryopreserved PBMCs from patients or healthy donors were initially stained with allophycocyanin-conjugated anti-CD25, FITC-conjugated anti-CD4 or anti-CD8, and PerCP-conjugated anti-CD3 antibodies, followed by intracellular staining for Foxp3 protein. The dot plots were gated on CD3⁺CD4⁺ T cells. The quadrants for Foxp3 were based on the isotype control antibody. This result is representative of 6 patients and 3 healthy donors.

Administration of high-dose IL-2 alters lymphocyte counts. (A) Patients received 3 daily doses of high-dose IL-2 as tolerated (total of 7-11 doses per patient). Patients were leukapheresed prior to the start of treatment on day 0 (PRE), during the peak of lymphocytosis or rebound (REB), and about 2 to 3 weeks after therapy (POST), as indicated by the arrows. Absolute lymphocyte counts (ALCs) were measured daily. The percent of CD4⁺ (B) and CD8⁺ (C) T cells per CD3⁺ T-cell population was quantified for 8 patients by staining PBMCs with anti-CD4 and anti-CD3 antibodies. The dot plots were gated on live (propidium idodide negative, PI^-) CD3⁺ T cells. For CD8 T cells, CD3⁺CD4^- T cells were considered as CD8 T cells in some experiments. Each symbol represents one patient. Leukaphereses samples for POST were available only for 5 patients. P values were determined using paired t test and adjusted for multiple comparisons.

Induction of CD25 expression by T cells following IL-2 administration. Cryopreserved PBMCs were triple-stained with allophycocyanin-conjugated anti-CD4, PE-conjugated anti-CD25, and FITC-conjugated anti-CD3 antibodies. (A) The dot plots were gated on PI^-CD3⁺ T cells. The gate for CD4⁺CD25^hi T cells was drawn based on the top 4% of CD4⁺ T cells expressing high levels of CD25 in the peripheral blood of each patient prior to IL-2 therapy (PRE). The same gate was used for rebound and post-treatment samples for each patient. The number represents the percentage of CD4⁺CD25^hi T cells per total CD4 T cells. The gate for CD8⁺CD25⁺ (CD4^-CD3⁺) was based on the isotype control antibody, and the number represents the percentage of CD8⁺CD25⁺ T cells per total CD8 T cells. The percent induction of CD25 expression of CD4 (B) and CD8 (C) T cells in 8 patients were quantified by using the gate for CD4⁺CD25^hi and for CD8⁺CD25⁺ T cells, respectively. To enhance for consistency and accuracy, PBMCs from PRE, REB, and POST for each patient were stained and analyzed at the same time. Leukaphereses samples for POST were available only for 5 patients. Each symbol represents one patient. The P values were calculated using paired t test and adjusted for multiple comparisons.

CD4⁺CD25^hi T cells from rebound phase were suppressive in vitro. To fractionate CD4⁺CD25^hi and CD4⁺CD25^- T cells, purified CD4 T cells isolated from rebound were stained with anti-CD25 PE and subsequently with anti-PE MicroBeads and isolated over magnetic columns as described in “Patients and methods.” We developed and optimized this procedure to consistently enrich for CD4⁺CD25^hi T cells without FACS sorting. (A) The dot plots were gated on CD3⁺CD4⁺ T cells and represent the initial starting CD4 T-cell population (Unfractionated) and the subsequent purified CD4CD25 subsets. The numbers represent the percentage of CD4⁺CD25^hi T cells for each fraction. (B, C) Freshly isolated CD4⁺CD25^- T cells (50 000 cells/well) were cocultured alone (CD25^-) or with CD4⁺CD25^hi T cells at different ratios and stimulated with anti-CD3 and irradiated T-depleted PBMCs (250 000 cells/well). Proliferation was assessed by [³H]thymidine incorporation pulsed on day 4 and harvested 18 hours later. The results represent the average [³H]thymidine incorporation (CPM) from 5 replicate wells per culture with calculated standard error of the mean. The P values were calculated using 2-sample t test. (D) CD4 T cells were initially purified from cryopreserved PBMCs collected during rebound from another patient, and stained with PE-conjugated anti-CD25, allophycocyanin-conjugated anti-CD4, and FITC-conjugated anti-CD3 antibodies, and subsequently separated into CD4⁺CD25^hi and CD4⁺CD25^- subsets using FACSVantage DiVa sorter. The result is the mean of [³H]thymidine incorporation in triplicate cultures with standard error of the mean, pulsed on day 6 and harvested 18 hours later.