Characterization of CD4⁺ T cell subsets. (A) PBMCs from an RCC patient were stained with anti-CD4/CD25 mAbs and analyzed by FACS. (B) CD4⁺CD25^neg, CD4⁺CD25^int (R1), and CD4⁺CD25^high (R2) T cells were sorted. For functional analysis, CD4⁺CD25^neg (left panel) and CD4⁺CD25^int (middle panel) T cells were stimulated with tetanus toxoid–loaded DCs (tetanus), DCs transfected with autologous RCC RNA (RCC), or with DCs transfected with autologous RE RNA (RE) at the indicated stimulator to responder ratios. After 48 hours, cells were pulsed with ³H-thymidine, and incorporation was determined using a liquid scintillation counter. CD4⁺CD25^high cells (right panel) were functionally validated by MLR. Mixture ratios of 1 CD4⁺CD25^high cell per T cell (Treg 1:1) or 1 CD4⁺CD25^high cell per 5 T cells (Treg 1:5) were added to the reaction, and inhibition of cell proliferation was analyzed. As a negative control, proliferation of CD4⁺CD25^high cells was determined in the presence of allogeneic DCs only (DC+Treg). Results are presented as means with SD calculated from triplicate wells. (C) FACS-based detection of GITR, CTLA-4, and FoxP3 by CD4⁺CD25^neg/int and CD4⁺CD25^high T cells subsets with or without stimulation using anti-CD3/CD28 mAb. Gray histograms represent isotypic controls. (D) Left panel: analysis of FoxP3 transcripts was performed by real-time PCR on indicated T cell populations. FoxP3 mRNA copy numbers were normalized to 1 × 10⁷ copies of β-actin mRNA. A representative result from 3 subjects is shown. Right panel: FoxP3 mRNA was amplified from CD4⁺ T cells isolated from RCC patients (n = 5) and healthy donors (n = 5). Differences in FoxP3 mRNA expression among groups were significant (P = 0.009).

Enhancement of T cell responses after Treg depletion. (A and B) CD4⁺CD25^high cells were isolated by FACS sorting and incubated for 6 hours in the presence of increasing concentrations of DAB₃₈₉IL-2 (left panel). In order to determine DAB₃₈₉IL-2–mediated toxicity, PBMCs and PBMCs admixed with CD4⁺/CD25^high cells at a 1:1 ratio were incubated with or without DAB₃₈₉IL-2 (5 nM) for 6 hours. In all experiments, cell viability was determined through MTT assays (A) or 7-AAD staining (B). Representative results from 3 evaluable subjects are presented. (C) Treg-depleted PBMCs (PBMC+DAB) or nondepleted PBMCs (PBMC–DAB) from an RCC patient were analyzed in allogeneic MLRs using DCs at a responder to stimulator ratio of 20:1. Cell proliferation was significantly inhibited when isolated CD4⁺/CD25^high cells were added to PBMCs at a 1:1 PBMC/CD4⁺CD25^high cell ratio (DC+Treg). This inhibition was reversible when the added CD4⁺/CD25^high cells were pretreated with DAB₃₈₉IL-2 (5 nM) for 48 hours (DC+Treg+DAB). Exposure of PBMCs to DAB₃₈₉IL-2 during the T cell–priming phase (day 2) led to complete inhibition of T cell proliferation (DC+DAB). (D) DCs transfected with mRNA encoding hTERT or MART-1 were used to stimulate CTL from Treg-depleted (filled symbols) or nondepleted (open symbols) human PBMCs.In addition, DCs loaded with MART-1–derived peptide 26-35 ELAGIGILTV (MART-1 pep) were used as stimulators. Following 2 stimulation cycles, CTLs were analyzed for their capacity to lyse their cognate (squares) or control targets (circles). As control targets, DCs loaded with GFP mRNA (mock transfected) or irrelevant peptide were used.

Depletion of Tregs in study subjects. (A) CD4⁺ T cells isolated from all DAB₃₈₉IL-2–treated study patients were analyzed by flow cytometry for expression of CD25 prior to and 4 days after intravenous administration; percentages of CD4⁺/CD25^high T cells are shown. (B) Reduction of FoxP3 mRNA copy numbers before and after DAB₃₈₉IL-2 treatment was determined by CD4⁺ T cells derived from 4 study subjects, as described in the legend to Figure 1D. The average FoxP3 mRNA copy number averaged from 5 healthy volunteers was used as control (C). (C) Functional analysis of Tregs isolated from study subjects prior to and after DAB386IL-2 administration. CD4⁺/CD25⁺ and CD4⁺/CD25^– T cell subsets were isolated from PBMC samples by magnetic bead separation, and Treg-mediated inhibition of activated CD4⁺/CD25^– indicator T cells was measured according to a protocol described previously (28).

Specificity of Treg depletion. (A) Calculated CD4⁺/CD25^high Treg frequencies in 2 study subjects (01-RCC-DAB; 03-RCC-DAB) prior to, 4 days after, and 2 weeks after final vaccination (study week 8; d 56). (B) IFN-γ ELISPOT were performed on sorted CD4⁺CD25^neg, CD4⁺CD25^int, and CD4⁺CD25^high T cell subsets using tetanus toxoid and CMV lysate–pulsed DCs as stimulators. (C–E) In separate experiments, IFN-γ ELISPOT and antigen-specific proliferation analyses were performed on T cells isolated prior to vaccination and 4 and 28 days after DAB₃₈₉IL-2 treatment (results from patient RCC-01-DAB). For ELISPOT assays, 1 × 10⁵ purified CD4⁺ T cells (C) or 1 × 10⁵ purified CD8⁺ T cells (D) were stimulated with 1 × 10⁴ DCs that were transfected with fluM1 mRNA, autologous PBMC RNA, CMV lysate (20 μg/ml), or tetanus toxoid (0.5 μg/ml). After 18 hours, visible spots were enumerated using an automated ELISPOT reader. The same stimulators and RCC RNA-transfected DCs were used in proliferation assays (E). For proliferation assays, isolated CD3⁺ T cells were used as responders. Assays were performed at a stimulator/responder ratio of 1:10. After 4 days, cells were pulsed with 1 μCi of ³H-thymidine, and incorporated radioactivity was determined after 16 hours by liquid scintillation counting.

In vivo induction of tumor-specific T cell responses. (A) CD8⁺ and CD4⁺ T cells were isolated from prevaccination (white bars) and postvaccination (black bars) PBMCs of patients treated with DAB₃₈₉IL-2 and RCC RNA-transfected DCs. For vaccination, 3 doses of 1 × 10⁷ cells injected intradermally every other week were administered. Isolated CD8⁺ and CD4⁺ T cells were stimulated for 18 hours with tumor RNA-transfected DCs, RE, or PBMC RNA-transfected DCs (controls). Visible spots were enumerated, and antigen-specific T cell frequencies were expressed as the number of spots forming cells per 1 × 10⁵ T cells. (B) Left panels: stimulation of tumor-specific CD8⁺ in 4 subjects treated with tumor RNA-transfected DCs alone. Right panels: summary of CD8⁺ and CD4⁺ T cell responses from 4 subjects receiving immunization alone (–DAB) or from 7 patients treated with combined therapy (+DAB). Bars indicate the median value of all subjects analyzed. Filled triangles represent T cell frequencies of individual patients. (C) Temporal evolution of tumor-specific CD8⁺ T cells after vaccination. IFN-γ ELISPOT analyses on sorted CD8⁺ T cells were performed as described in A. Frequencies of tumor-specific T cells prior to, during, and after immunization are presented for 2 patients who received 3 vaccinations with tumor mRNA-transfected DCs alone (11-RCC) or were treated with DAB₃₈₉IL-2 (01-RCC-DAB) followed by vaccination.

In vivo induction and cytokine profile of RCC-specific CD4⁺ T cell responses. (A) CD4⁺ T cells were isolated from pre- (white bars) and post-vaccination (black bars) PBMC samples of 3 study subjects (representative data from patient RCC-01-DAB are shown) who received DAB₃₈₉IL-2 (18 μg/kg) followed by vaccination with RCC RNA-transfected DCs (2 cycles of 1 × 10⁷ cells per treatment). Cells were stimulated for 18 hours with autologous PBMC RNA-, RE RNA-, or RCC RNA-transfected DCs. IFN-γ (left panel) or IL-4–expressing T cells (right panel) were enumerated using an automated ELISPOT reader, and antigen-specific T cell frequencies were expressed as the number of spot-forming cells per 1 × 10⁵ CD4⁺ T cells. Staphylococcal enterotoxin B (SEB) at a concentration of 10 μg/ml was used as a positive control in the IL-4 ELISPOT assays (right panel). (B) The cytokine expression profile of CD4⁺ T cells prior to (gray) and after (white) vaccination was measured after overnight (18 hours) stimulation with RCC (DC+RCC) or RE RNA-transfected DCs (DC+RE) using human Th1/Th2 cytometric bead arrays. Culture supernatants were used to determine expression of the Th-1 cytokines IFN-γ, TNF-α, and IL-2 as well as the Th-2 type cytokines IL-4 and IL-10.