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Vaccination of patients with dendritic cell (DC)/breast carcinoma fusions stimulated antitumor immune responses in a majority of patients with metastatic disease but only a subset demonstrate evidence of tumor regression. To define the factors that limit vaccine efficacy, we examined the biological characteristics of DC/breast carcinoma fusions as APCs and the nature of the vaccine-mediated T cell response. We demonstrate that fusion of DCs with breast carcinoma cells up-regulates expression of costimulatory and maturation markers and results in high levels of expression of IL-12 consistent with their role as activated APCs. Fusion cells also express the chemokine receptor CCR7, consistent with their ability to migrate to the draining lymph node. However, DC/breast cancer fusions stimulate a mixed T cell response characterized by the expansion of both activated and regulatory T cell populations, the latter of which is characterized by expression of CTLA-4, FOXP3, IL-10, and the suppression of T cell responses. Our results demonstrate that IL-12, IL-18, and TLR 9 agonist CpG oligodeoxynucleotides reduce the level of fusion-mediated regulatory T cell expansion. Our results also demonstrate that sequential stimulation with DC/breast carcinoma fusions and anti-CD3/CD28 results in the marked expansion of activated tumor-specific T cells. These findings suggest that DC/breast carcinoma fusions are effective APCs, but stimulate inhibitory T cells that limit vaccine efficacy. In contrast, exposure to TLR agonists, stimulatory cytokines, and anti-CD3/CD28 enhances vaccine efficacy by limiting the regulatory T cell response and promoting expansion of activated effector cells.
Breast cancer cells express unique Ags that are recognized by the host immune repertoire and serve as potential targets for tumor immunotherapy. However, tumor cells evade host immunity, in part, by presenting Ag in the absence of costimulation and the suppression of native APCs (1). Dendritic cells (DCs)3 represent a complex network of APCs that are primarily responsible for initiation of primary immunity and modulation of the immune response (2, 3). Partially mature DCs are located at sites of Ag capture, excel at the internalization and processing of exogenous Ags, but are poor stimulators of T cell responses. Upon activation, DCs undergo maturation characterized by the increased expression of costimulatory molecules and CCR7, the chemokine receptor which promotes migration to sites of T cell traffic in the draining lymph nodes (4). Presentation of Ag by immature DCs may induce T cell tolerance (5). Breast carcinomas inhibit DC development through the secretion of IL-10, TGF-β, and vascular endothelial growth factor, resulting in the accumulation of immature DCs in the tumor bed that potentially suppress antitumor responses (6-9). Conversely, activated DCs can be generated by cytokine-mediated differentiation of DC progenitors ex vivo. DC maturation and function can be further enhanced by exposure to CpG oligodeoxynucleotides (ODN) (10, 11).
We have developed a cancer vaccine in which patient-derived tumor cells are fused with autologous DCs generated ex vivo. DC/tumor fusions express a broad array of tumor Ags presented in the context of DC-mediated costimulation. In animal models, vaccination with DC/tumor fusions protects against an otherwise lethal challenge of tumor cells and effectively eradicates established disease (12-15). Fusions of patient-derived breast carcinoma cells and DC stimulate T cell-mediated lysis of autologous tumor cells in vitro (16). In a clinical trial for patients with metastatic breast carcinoma, vaccination with autologous DC/tumor fusions induced antitumor immunity in a majority of patients while clinical responses were observed in a only subset of patients (17).
An effective cancer vaccine must have the capacity to present tumor Ags in the context of stimulatory signaling, migrate to sites of T cell traffic, and induce the expansion of activated effector cells with the ability to lyse tumor targets. One concern regarding the DC/breast carcinoma fusions is that tumor cells in the vaccine preparation may inhibit its function as an APC. Another potential issue limiting response to vaccination is the presence of regulatory T cells that suppress T cell activation (18-20). Regulatory T cells deliver inhibitory signals via direct cell contact and the release of cytokines that play a role in mediating tumor-associated anergy. Regulatory T cells are increased in the circulation, tumor bed, and lymph nodes of patients with malignancy and their presence has been associated with worse outcomes (21-23). Paradoxically, studies have demonstrated that vaccination may lead to the expansion of regulatory T cells that ultimately blunt response. In animal models, depletion of regulatory T cells resulted in enhanced response to tumor vaccines (24, 25). Several strategies have been examined to enhance vaccine efficacy and promote T cell polarization toward an activated phenotype. Activation of innate immunity through ligation of TLR 9 potently stimulates T cell responses (26-31). Similarly, exposure to stimulatory cytokines, such as IL-12 and IL-18, results in T cell polarization toward a Th1 phenotype (14, 32-42). Ligation of the T cell/costimulatory complex (CD3/CD28) has also been shown to promote the activation of T cells when administered in the context of other stimulatory signals (43-46).
In the present study, we examined the phenotypic characteristics of DC/breast carcinoma fusions with respect to their function as APCs. We demonstrate that DC/breast carcinoma fusions exhibit features consistent with activated DCs as manifested by high levels of expression of costimulatory molecules, IL-12, and the chemokine receptor CCR7. In fact, immature DCs undergo maturation following polyethylene glycol (PEG)-mediated fusion with breast carcinoma cells. However, we also demonstrate that DC/tumor fusions stimulate a mixed response of activated and regulatory T cells, the latter of which potentially interfere with the development of antitumor immunity. We show that DC/breast carcinoma fusion cells induce the expansion of CD4+CD25+high T cells, which uniformly express FOXP3 and are potent inhibitors of CD4+CD25− T cells in a coculture system in vitro. Additionally, we also investigated whether the administration of a second stimulatory signal would favor the development of an activated antitumor immune response. We show that the addition of IL-12, IL-18, and the TLR 9 agonist CpG ODN decreases fusion-mediated expansion of regulatory T cells. Most notably, combined stimulation with DC/breast carcinoma fusions and anti-CD3/CD28 results in the dramatic expansion of tumor-specific T cells with an activated phenotype.
PBMCs were isolated from leukopaks from normal donors and from peripheral venous blood collected from patients with breast cancer as per an institutionally approved protocol. PBMCs underwent Histopaque −1077 (Sigma-Aldrich) density gradient centrifugation and were plated in tissue culture flasks (BD Biosciences) in RPMI 1640 culture media containing 2 mM l-glutamine (Mediatech) and supplemented with heat-inactivated 10% human AB male serum (Sigma-Aldrich), 100 U/ml penicillin, and 100 μg/ml streptomycin (Mediatech) (complete medium) for 2 h at 37°C in a humidified 5% CO2 incubator. The monocyte-enriched adherent fraction was cultured in complete medium containing GM-CSF (1000 U/ml) (Berlex) and IL-4 (1000 U/ml) (R&D Systems) for 5 days to generate immature DCs. A fraction of the DC preparation underwent further maturation by culturing the cells for an additional 48 h in the presence of TNF-α (25 ηg/ml) (R&D Systems) or the combination of TNF-α (25 ηg/ml), IL-1β (10 ηg/ml), IL-6 (1000 U/ml) (R&D Systems), and PGE2 (1 μg/ml) (Calbiochem).
T cells were isolated from the nonadherent PBMC fraction using a T cell enrichment column (R&D Systems) or nylon wool column (Polysciences). Purity of T cells by both methods was >90% as determined by FACS analysis of CD3 surface expression. T cells were classified as allogeneic when derived from a third party donor and autologous when derived from the same donor from whom the DC fusion partner was derived.
Primary breast carcinoma cells were obtained from malignant effusions or resected tumor lesions as per an institutionally approved protocol. Human breast carcinoma cell lines MCF-7 and ZR-751 were purchased from ATCC. All tumor cell lines were maintained in DMEM (high glucose) or RPMI 1640 supplemented with 2 mM l-glutamine, 100 U/ml penicillin, 100 μg/ml streptomycin, and 10% heat-inactivated FBS (HyClone).
DC/breast carcinoma fusions were prepared as previously described (16). Tumor cells were mixed with immature or mature DC preparations at ratios of 1:3-1:10 (dependent on cell yields) and washed in serum-free prewarmed RPMI 1640 culture media. The cell pellet was resuspended in 50% PEG solution (molecular mass: 1450)/DMSO solution (Sigma-Aldrich). After 3 min at room temperature, the PEG solution was progressively diluted with prewarmed serum-free RPMI medium and washed twice with serum free media. The fusion preparation was cultured for 5–7 days in 5% CO2 at 37°C in complete medium with GM-CSF (500 IU/ml). Fusion cells were isolated for subsequent analyses by FACS gating around the population that coexpressed unique DC and tumor Ags as outlined below.
DCs and breast carcinoma cells were incubated with primary mouse anti-human mAbs directed against HLA-DR, CD11c, CD14, CD80, CD86, CD83, CD40, CD54, MUC1, cytokeratin (CT), and matching isotype controls (BD Pharmingen), washed, and cultured with FITC-conjugated goat anti-mouse IgG1 (Chemicon International). Cells were fixed in 2% paraformaldehyde (Sigma-Aldrich) and underwent flow cytometric analysis using FACScan (BD Biosciences) and CellQuest Pro software (BD Biosciences). DC/breast carcinoma fusions preparations were subjected to dual staining to quantify the percentage of cells that coexpressed unique DC (CD11c-Cychrome) and tumor Ags (MUC1 or CT-FITC). Fusion cells were isolated by FACS gating and stained with PE-conjugated mouse anti-human Abs directed against CCR7, CD80, CD86, or CD83. The percentage of fusion cells expressing these markers was determined by multi-channel flow cytometric analysis. Alternatively, an aliquot of fusion cells was pulsed with GolgiStop (1 μg/ml; BD Pharmingen), permeabilized by incubation in Cytofix/Cytoperm plus (containing formaldehyde and saponin; BD Pharmingen), and washed in Perm/Wash solution (BD Pharmingen). The cells were then incubated with PE-conjugated anti-human IL-10 or IL-12 (Caltag Laboratories) or a matched isotype control Ab for 30 min, washed twice in Perm/Wash solution, and fixed in 2% paraformaldehyde (Sigma-Aldrich). The IL-12 Ab recognizes the IL-12 p40 monomers, homodimers, and the p70 heterodimers, but not the p35 subunit. A minimum of 1 × 104 events were acquired for analysis. To determine whether DC/breast carcinoma fusion cells coexpressed IL-10 and IL-12, fusion cells were labeled with PE Cy7-conjugated HLA-DR (eBioscience) and Cy5-conjugated DF3 Ab (DF3-conjugated using the Cy5 reactive Dye Kit; Amersham Biosciences) followed by intracellular staining with IL-10 (PE-conjugated) and IL-12 (FITC-conjugated) or matched isotype controls. The fusion cells were analyzed by 4-color flow cytometry using the BD LSR II analyzer (BD Biosciences) and data was analyzed using the BD FACSDiva software (BD Biosciences).
Approximately 1.2 × 104 cells were spun onto slides (Cytospin; Thermo Shandon), allowed to dry, and fixed with acetone. The slides were incubated with primary mouse anti-human mAbs MUC1 and CT and an isotype-matched negative control at room temperature for 1 h, washed, incubated with 1:100 biotinylated F(ab’)2 of horse anti-mouse IgG (Vector Laboratories), washed, and incubated for 30 min with avidin-biotin complex reagent solutions (Vector Laboratories) followed by 3 amino-9-ethyl carbazole solution (Vector Laboratories). Cells were then stained for HLA-DR, CD86, or CD83 with the avidin-biotin complex-alkaline phosphatase kit (Vector Laboratories). Slides were washed, fixed in 2% paraformaldehyde (Sigma-Aldrich), and analyzed using an Olympus AX70 microscope.
To assess their capacity to stimulate allogeneic T cell proliferation, immature and mature DCs and DC/breast carcinoma fusion cell preparations were cocultured with allogeneic normal donor-derived T cells at a ratio of 1:10, 1:30, 1:100, 1:300, and 1:1000 in 96-well U-bottom culture plates (Costar) for 5 days at 37°C and 5% CO2. T cell proliferation was determined by incorporation of [3H]thymidine (1μCi/well; 37kBq; NEN-DuPont) added to each well 18 h before the end of the culture period. There-after, the cells were harvested onto glass fiber filter paper (Wallac) using an automated TOMTEC harvester (Mach II), dried, placed, and sealed in BetaPlate sample bag (Wallac) with 10 ml of ScintiVerse (Fisher Scientific). Cell bound radioactivity was counted in a liquid scintillation counter (Wallac; 1205 Betaplate). Data are expressed as stimulation index (SI). The SI was determined by calculating the ratio of [3H]thymidine incorporation (mean of triplicates) over background [3H]thymidine incorporation (mean of triplicates) of the unstimulated T cell population.
The profile of secreted cytokines by T cells cultured with immature and mature DC/breast carcinoma fusions was determined using the cytometric bead array (CBA) kits (BD Biosciences). Supernatants from unstimulated T cells or cells exposed to unfused DC and breast carcinoma served as controls. Supernatants were collected before cell harvest and frozen at −80°C. Concentrations of IL-2, IL-4, IL-5, IL-10, IFN-γ, TNF-α, IL-12, IL-6, IL-1β, and IL-8 were quantified using an inflammatory CBA kit as per standard protocol. Briefly, the kits provided a mixture of six microbead populations with distinct fluorescent intensities (FL-3) that are precoated with capture Abs specific for each cytokine. Culture supernatant or the provided standardized cytokine preparations were added to the premixed microbeads and then cultured with secondary PE-conjugated Abs. Individual cytokine concentrations were indicated by their fluorescent intensities (FL-2) and then computed using the standard reference curve of Cellquest and CBA software (BD Pharmingen). Interassay reproducibility was assessed using two replicate samples of three different levels of the human standards in three separate experiments.
DC/breast carcinoma fusion cell preparations generated with immature and mature DCs were cocultured with autologous T cells at a ratio of 1:10 for 7–10 days. DC/breast carcinoma fusions generated with DC autologous to T cell effectors were used as target cells in a standard 5-h 51Cr-release assay. Target cells (2 × 104 cells/well) were incubated with 51Chromium (NEN-DuPont) for 1 h at 37°C followed by repeated washes. 51Cr release was quantified following 5-h coculture of effector and target cell populations at a ratio of 30:1 or 10:1. Percentage cytotoxicity was calculated using mean of triplicates by a standard assay as follows: % specific cytotoxicity = [(sample counts − spontaneous counts)/(maximum counts − spontaneous counts)] × 100. Spontaneous release was <25% of the maximum 51Cr uptake. As a control, lysis of targets by unstimulated T cells and T cells stimulated by unfused DCs were assessed.
Ag-specific MUC1+CD8+ T cells were identified using PE-labeled HLA-A*0201+ iTAg MHC class I human tetramer complexes (Beckman Coulter) composed of four HLA MHC class 1 molecules each bound to MUC1-specific epitopes M1.2 (MUC112–20) LLLLTVLTV (47). An A*0201 irrelevant peptide MHC class I tetramer with no known specificity was provided by Beckman Coulter as a negative control. Nonadherent cells were cocultured with DC/breast carcinoma fusion cells for 5 days, harvested, incubated with the MUC1 or control tetramer, and then stained with FITC-conjugated CD8 Ab. Cells were washed and analyzed by bidimensional FACS analysis. A total of 3 × 105 events were collected for final analysis. Similarly, nonadherent unstimulated cells were analyzed in parallel.
Autologous and allogeneic T cell preparations were cocultured with mature DC/breast carcinoma fusions for 5 days at a 10:1 ratio. The cell preparations were incubated with FITC-conjugated anti-CD4, Cychrome-conjugated anti-CD25, and PE-conjugated anti-CD69, antiglucocorticoid-induced TNF receptor (GITR), or anti-CTLA-4. Alternatively, cells were permeabilized and cultured with PE-conjugated Ab directed against IFN-γ, IL-10, IL-4, or FOXP3. Cells were subsequently analyzed by multi-channel flow cytometry. In some studies, CD4+ T cells were isolated by magnetic microbead isolation (Miltenyi Biotec), and the resultant population were subjected to a two staining procedure with anti-CD25 Ab and the indicated marker.
In an effort to segregate activated and regulatory T cell populations based on the expression of CD4+CD25−, CD4+CD25+low and CD4+CD25+high, 40–50 × 106 of the CD4+ T cells were positively selected (>97% purity) from resting and fusion-stimulated T cell populations and sorted using a BD FACSAria cell sorting instrument (BD Biosciences). The cells were incubated with anti-CD4 TC (IgG2a; Invitrogen) and anti-CD25-FITC (IgG1; BD Pharmingen) and separated into CD4+CD25−, CD4+CD25+low, and CD4+CD25+high fractions by FACS sorting as previously described (18). Intracellular staining for FOXP3 was performed on cells directly after sorting using the PCH101 anti-FOXP3 Ab (eBioscience) per the manufacturer’s instructions. CD4+CD25− T cells (5 × 104 cells/well) were cocultured in triplicate with equal numbers of CD4+D25+low, CD4+D25+high, or CD4+CD25− T cells (as controls) in a 1:1 ratio, in the presence of irradiated (3500 Rads) autologous T cell depleted PBMCs as a source of APCs. The cultures were pulsed with tetanus toxoid (10 μg/ml), anti-CD3 Ab (1 μg/ml) (clone UCHT1; BD Pharmingen), or PHA (4 μg/ml) (without autologous PBMCs). T cell proliferation was quantified after 4 days of culture period by uptake of [3H]thymidine (1 μCi [0.037 MBq] per well) following overnight pulsing.
DC/breast carcinoma fusions were cocultured for 5–7 days with autologous T cells in the presence or absence of IL-12 (10 ng/ml; R&D Systems), IL-18 (10 ng/ml; R&D Systems), or CpG ODN (10 μg/ml; Coley Pharmaceutical Group). The recombinant human IL-12 added to the cultures was the p70 heterodimeric cytokine (R&D Systems). The CpG ODN (C-2395) consisted of a hexameric CpG motif, 5′-TCGTCGTTTT-3′, linked by a T spacer to the GC-rich palindrome sequence 5′-CGGCGCGCGCCG-3′ (48). A control CpG ODN (class B-2137) without stimulatory sequences was simultaneously tested in parallel in each experiment. Regulatory and activated T cell populations were quantified as outlined above.
T cells were activated for 48 h by exposure to the immobilized mAbs, anti-CD3 (clone-UCHT1; BD Pharmingen), and anti-CD28 (clone-CD28.2; BD Pharmingen; CD3i/CD28i). Twenty-four-well non-tissue culture-treated plates (Falcon) were coated with each of the Abs (1 μg/ml in PBS) at 0.5 ml/well and left overnight at 4°C. The plates were blocked with 1% BSA and T cell preparations were added at a density of 2 × 106 cells/well. T cells were stimulated with anti-CD3/CD28 (48 h) or DC/breast carcinoma fusions alone (5–7 days), fusions followed by exposure to anti-CD3/CD28, or anti-CD3/CD28 followed by fusion cells. T cells were harvested and proliferation was determined by uptake of tritiated thymidine. T cells binding the MUC1 tetramer were quantified by FACS analysis as outlined above. The percentage of T cells expressing markers consistent with a regulatory (FOXP3) and activated (CD69, IFN-γ) phenotype were quantified.
Results are expressed as mean ± SEM. For comparisons, Student’s t test was used and values of p < 0.05 were considered as significant.
Tumor cells suppress host immunity, in part, by disrupting the development and function of APCs. A potential issue concerning the effectiveness of the DC/tumor fusion vaccine is whether the tumor cell fusion partner will inhibit DC differentiation and interfere with Ag presentation by the fusion vaccine. To assess this question, we examined the phenotypic and functional characteristics of fusions generated with immature and mature DCs. Immature and mature DCs were generated from patients with breast cancer and from leukopak preparations acquired from volunteer donors. Adherent PBMC were cultured for 1 wk with GM-CSF and IL-4 to generate partially mature DCs. Maturation was induced by exposure to TNF-α for 48–96 h. Both immature and mature DC preparations strongly expressed the costimulatory molecule CD86, [75% (range: 25–98%) and 84% (45–99%), respectively] and exhibited low levels of CD14 expression (n = 15) (Fig. 1A). However, mature DC demonstrated a statistically significant increase in mean expression of CD80 [20% (3–65%) vs 9% (2–46%); p = 0.05] and CD83 [31% (3–77%) vs 7% (3–15%); p = 0.0003]. Similar phenotypic changes were observed following DC maturation with CD40L (data not shown). As a measure of their functional capacity as APCs, DC preparations were examined for their ability to stimulate allogeneic T cell proliferation. Mature as compared with immature DCs stimulated higher levels of allogeneic T cell proliferation (data not shown).
We subsequently examined the phenotypic characteristics of the fusion cell populations generated with immature and mature DC. Immature and mature DC populations were fused with primary patient-derived breast cancer cells or MCF-7 human breast carcinoma cells by coculture with PEG. Fusion cells were quantified by determining the percentage of cells that coexpressed unique tumor (MUC1 and/or CT) and DC (CD11c) Ags (Fig. 1B). Equivalent mean fusion efficiencies were observed following fusion of tumor cells with mature (11% ± 1.6 SEM) and immature (7% ± 1.2 SEM) DC (n = 12). Using primary tumor cells obtained from patient derived samples, fusion cells were isolated by FACS gating of cells that coexpressed DC and tumor-derived Ags (Fig. 1B). Expression of CD86 was uniformly observed in both immature DC/breast carcinoma (89%) and mature DC/breast carcinoma (82%) fusion populations (n = 6) (Fig. 1C). The maturation marker, CD83, was detected in 46 and 51% of immature and mature fusion cell populations, respectively (p = 0.5, NS) (Fig. 1, B and C). A significant increase in CD83 and CD86 expression was observed in immature DC/breast carcinoma fusions as compared with immature DCs. Immunocytochemical staining demonstrated expression of DR, CD86, and CD83 by the immature DC/tumor fusions (Fig. 1, D–G). These studies demonstrate that fusion of DC with breast carcinoma cells results in phenotypic characteristics consistent with maturation and activation, and not inhibition of DC differentiation.
As a measure of their potency as APCs and their capacity to stimulate Th1 responses, we next examined expression of IL-12 and IL-10 by the fusion cell populations generated with primary tumor cells obtained from patient samples (Fig. 2, A and B). Fusion cells were isolated by FACS gating of cells that coexpressed DC and tumor-derived Ags. The mean percentage of fusion cells that express IL-12 and IL-10 did not differ between the fusion cell populations. IL-12 was expressed by ~40 (±6.7 SEM) and 49% (±6.3 SEM) (p = 0.35, NS) and IL-10 by ~36.3 (±6.4 SEM) and 40% (±6.4 SEM; n = 11) (p = NS) of the immature and mature DC/breast carcinoma fusions, respectively (n = 12). We subsequently analyzed whether IL-12 and IL-10 was expressed by distinct populations of DC/breast carcinoma fusions. Immature and mature DC/breast carcinoma fusions were isolated by FACS gating of cells that coexpressed HLA-DR and MUC1. The gated cells underwent intracellular FACS analysis for IL-12 and IL-10 expression. IL-12 and IL-10 were expressed by a single DC/breast carcinoma fusion population (Fig. 2, C and D).
The chemokine receptor, CCR7, directs cell migration to sites of T cell traffic in the draining lymph nodes and is characteristically expressed by DCs that are undergoing maturation and activation. As a measure of their migratory capacity, expression of CCR7 was determined for fusions generated with primary breast tumors and immature or mature DC (Fig. 2E). CCR7 was prominently expressed on both immature and mature fusion populations, suggesting that tumor-DC fusion resulted in the expression of a mature and activated phenotype. Mean CCR7 expression was observed in 33 (±9 SEM) and 38% (±7.3 SEM; n = 11) of the immature and mature DC/breast cancer fusions, respectively (n = 12). In contrast, mean expression of CCR7 by immature DCs was 3.8%.
The functional capabilities of mature as compared with immature DC/tumor fusion preparations were analyzed by comparing their capacity to stimulate T cell proliferation and cytokine production. Fusion cell populations were cocultured with autologous T cells for 5 days and proliferation was determined by measuring uptake of tritiated thymidine after overnight pulsing (Fig. 2F). Proliferation was measured as the T cell SI (Stimulated T cells/Unstimulated T cells). Both immature and mature DC/breast cancer fusions stimulated autologous T cell proliferation with SI of 3.3 (±1.4 SEM; n = 6) and 3.5 (±1.4 SEM; n = 6), respectively. We also quantified cytokine secretion generated by coculture of DCs, immature or mature DC/primary breast carcinoma fusions with autologous T cell populations using the BD CBA system (BD Biosciences) (Fig. 3, A and B). Mean levels of IFN-γ following stimulation with immature and mature DC/breast cancer fusions were 2188 and 2252 pg/ml, respectively. These levels were significantly greater than those seen with T cells cultured with unfused autologous DC (685 pg/ml). In contrast, secretion of IL-12, IL-4, IL-10, IL-2, and TNF-α was not increased following stimulation with fusion cells.
Both immature and mature DC/tumor populations were capable of generating significant levels of target-specific killing, as demonstrated by the lysis of autologous tumor or semiautologous fusion targets. CTL activity did not differ between the fusion populations (n = 5; stimulated by DC/breast carcinoma fusions). Mean CTL lysis for effector:T cell ratio of 30:1 was 27 and 21% for T cells stimulated with immature and mature DC/breast cancer fusions, respectively (p = NS) (Fig. 4A). In contrast, only background levels of killing were observed following coculture of targets with unstimulated T cells or those stimulated with unfused DCs. To assess the capacity of the fusion vaccine to stimulate T cell responses directed against a specific tumor Ag, we assessed whether HLA-A2.1+ T cells stimulated by DC/breast carcinoma fusions recognized MUC1. Selective expansion of CD8+ T cells binding the MUC1 tetramer was observed following stimulation with fusions generated with DCs and primary breast carcinoma cells (Fig. 4B). In a single experiment, 23.6% of the CD8+MUC1 tetramer+ T cells were observed to express IFN-γ. In a series of experiments, the mean percentage of CD8+ T cells binding the MUC1 tetramer were 5.49% (±0.46 SEM; n = 3) as compared with 1.3% (±0.06 SEM; n = 3) of CD8+ T cells binding the negative tetramer (p = 0.005). These results indicate that DC/breast carcinoma fusions exhibit an activated phenotype with strong expression of costimulatory molecules, stimulatory cytokines, and chemokine receptors enabling them to migrate to sites of T activation. In addition, DC/breast carcinoma fusions stimulate antitumor CTL responses including the expansion of T cells targeting defined tumor Ags.
Having characterized DC/breast carcinoma fusions as potent APCs with the capacity to elicit T cell responses, we examined whether DC/tumor fusions induce the expansion of regulatory as compared with activated T cells. Stimulation with DC/breast carcinoma fusions using MCF-7 (n = 13) or primary breast carcimona cells (n = 7) did not result in an increase in the percentage of total CD4+CD25+ T cells (10% ± 1.3 SEM; n = 20) as compared with unstimulated T cells (8.3% ± 1.1 SEM; n = 20) (Fig. 5A). Note-worthy, higher levels of CD4+CD25+ cells were observed following stimulation with fusions generated with primary breast carcinoma cells as compared with the MCF-7 cell line. Although both activated memory effector cells and regulatory T cells coexpress CD4 and CD25, regulatory T cells may be differentiated by their relatively high level of CD25 expression and the presence of other markers such as GITR, CTLA-4, and FOXP3. In contrast, CD69 is characteristically expressed by activated T cells. Mature DCs were fused to a human breast carcinoma cell line (MCF-7) and cocultured with autologous or allogeneic T cells for 5 days. CD4+CD25+ cells were quantified by flow cytometric analysis and further characterized with respect to expression of cell surface markers and cytokine profile. CD4+ T cells were positively selected from this population using CD4+ magnetic beads. FACS analysis of the resultant CD4+ T cells demonstrated a purity of greater than 97%. However, coculture of fusion cells and autologous T cells resulted in a 6.3-fold increase in CD4+CD25+ T cells that expressed CD69, (4.7%-unstimulated T cells; 29.5-fusion stimulated cells, n = 5; p = 0.01) consistent with an activated phenotype (Fig. 5B). Stimulation with mature DC/breast carcinoma fusions also resulted in 9- and 5.2-fold increase in CD4+CD25+ T cells that expressed GITR and CTLA-4, respectively (Fig. 5B). These findings suggest that both activated and inhibitory T cell populations are expanded by DC/breast carcinoma fusions (Fig. 5B). Noteworthy, fusion stimulation of allogeneic T cells resulted in a similar increase in CD4+CD25+CD69+ T cells (5-fold), but a significantly greater expansion of GITR-(25-fold) and CTLA-4-(15-fold) positive populations (Fig. 5C).
We also assessed the profile of cytokine expression in the CD4+CD25+ T cell population following stimulation with DC/breast carcinoma (MCF-7 or ZR75) fusions using intracellular flow cytometric analysis. The mean percentage of CD4+CD25+ T cells expressing IFN-γ was 40 (±6.9 SEM) and 68% (±6.1 SEM) before and following fusion cell stimulation (p = 0.005), respectively (n = 14) (Fig. 5D). Similarly, the percentage of CD4+CD25+ T cells expressing the inhibitory cytokine, IL-10 (Fig. 5D) rose from 20 (±4.9 SEM) to 59% (±8.4 SEM) (p = 0.0002). Finally, we assessed the impact of fusion cell stimulation on the intracellular expression of FOXP3, a marker considered to be specific for regulatory T cells. FOXP3 expression increased from 26.5 (±5.4 SEM; n = 9) to 63% (±10.6 SEM; n = 9) (p = 0.01) of the unstimulated and fusion stimulated CD4+CD25+ T cell populations, respectively (Fig. 5D). As such, fusion cells induce the expansion of both immunostimulatory and immunosuppressive elements resulting in a complex response in which regulatory T cells may prevent the development of sustained effective antitumor immunity.
To further define the functional characteristics of T cells stimulated by DC/breast carcinoma fusions, CD4+CD25−, CD4+CD25+low, and CD4+CD25+high T cells were separated by flow cytometric sorting (Fig. 6A). Fusion stimulation resulted in an increase in the CD4+CD25+low and CD4+CD25+high fractions. Consistent with a regulatory T cell phenotype, CD4+CD25+high T cells uniformly expressed FOXP3. In contrast, FOXP3 expression was seen in only a minority of CD4+CD25+low cells and was absent from CD4+CD25− cells. In a series of experiments, the mean percentage of FACS sorted CD4+CD25+high T cells that expressed FOXP3 were 86.9% (±7.1 SEM; n = 3) as compared with 11.73% (±2.6 SEM; n = 3) of CD4+CD25+low and 0.52% (±0.2 SEM; n = 3) of CD4+CD25−, respectively. To assess the functional properties of these populations, we examined their ability to suppress T cell proliferation in response to TCR ligation with anti-CD3 or exposure to the tetanus toxoid recall Ag. Consistent with regulatory T cell phenotype, addition of CD4+CD25+high cells to CD4+CD25− cells at a ratio of 1:1 significantly inhibited proliferative responses to anti-CD3 (7-fold decrease; p = 0.03) and tetanus toxoid (11.5-fold decrease; p = 0.0002) (Fig. 6B). Modest inhibition was also observed of PHA-mediated stimulation of CD4+CD25− cells. The degree of suppression correlated with the levels of CD4+CD25+high cells added to the culture (data not shown). In contrast, significant inhibition of T cell responses to anti-CD3, tetanus toxoid, and PHA was not observed following the addition of CD4+CD25+low or CD4+CD25− cells. These data demonstrate that DC/breast carcinoma fusion cells induce the expansion of distinct T cell populations with phenotypic and functional characteristics of regulatory and activated T cells, respectively.
In an effort to bias the T cell response toward an activated phenotype and limit the influence of regulatory T cells, we studied the effect of the TLR 9 agonist, CpG ODN on vaccine response. TLR agonists activate elements of the innate immune response and have been shown to augment vaccine efficacy. We examined the capacity of CpG ODN to modulate fusion-mediated stimulation of activated and inhibitory T cell populations by quantifying expression of IFN-γ as compared with IL-10 and FOXP3 in CD4+CD25+ cells. We also examined the effect of adding the stimulatory cytokines IL-12 and IL-18 on the phenotypic profile of T cells cocultured with DC/breast carcinoma fusions. A 2.5-fold increase was seen in the fusion stimulated CD4+CD25+ T cells in the presence of CpG ODN and IL-18, respectively (p = 0.0004 and p = 0.006). In contrast, no significant increase in CD4+CD25+ cells was observed when IL-12 was added to the cocultures of T cells and DC/breast cancer fusions (Fig. 7A).
The addition of CpG, IL-12, or IL-18 decreased the percentage of CD4+CD25+ with phenotypic characteristics of regulatory T cells as determined by FOXP3 expression (p = 0.024, p = 0.042, and p = 0.016, respectively). In concert with these findings, expression of IL-10 in the CD4+CD25+ T cells was significantly lowered in cocultures pulsed with CpG ODN (19.8% ±4.1 SEM, n = 7; p = 0.002) and IL-18 (18.3% ±5.1 SEM, n = 4; p = 0.0004) as compared with T cells stimulated by DC/breast carcinoma fusions alone (59.3% ±8.4 SEM, n = 14) (Fig. 7B). Notably, a decrease in the mean percentage of CD4+CD25+ T cells expressing IFN-γ was also seen following the addition of CpG and IL-18 to cocultures of fusions and autologous T cells (Fig. 7C). These results indicate that the addition of IL-12 or TLR agonists enhances vaccine efficacy by limiting the presence of immunosuppressive regulatory cells.
As another strategy to bias the vaccine response toward immune activation, we examined the effect of ligation of the TCR/costimulatory complex using Abs directed against CD3 and CD28. Exposure to anti-CD3/CD28 provides an Ag-independent stimulus resulting in the expansion of activated or inhibitory T cells, dependent on the nature of the surrounding immunologic milieu. We hypothesized that sequential stimulation with DC/breast carcinoma fusions followed by anti-CD3/CD28 would amplify the response of T cells that had been primarily activated by the fusion vaccine.
Limited proliferation of T cells was observed following exposure to anti-CD3/CD28 alone (SI: 1.5 ± 0.5 SEM; n = 7) or DC/breast carcinoma fusions (SI: 3.1 ± 1.2 SEM; n = 7) (Fig. 8A) However, a marked increase in T cell expansion was noted when T cells were first stimulated with DC/breast carcinoma fusions and then expanded with anti-CD3/CD28 (SI: 23 ± 8.73 SEM; n = 7). Noteworthy, no increase in proliferation was observed when T cells were first exposed to anti-CD3/CD28 and then cultured with DC/breast carcinoma fusions (SI: 1.6 ± 0.3 SEM; n = 6). Sequential stimulation with DC/breast carcinoma fusions generated with primary tumor cells and anti-CD3/CD28 resulted in the specific expansion of tumor reactive T cells. Exposure to anti-CD3/CD28 following fusion cell stimulation induced a 13.7 mean-fold increase in MUC1 tetramer binding cells (n = 3) (Fig. 8B). The percentage of MUC1 tetramer+ cells remained at baseline levels following stimulation with anti-CD3/CD28 alone. With regard to the phenotype of the expanded T cell population, the percentage of T cells expressing the CD4+CD25+ phenotype was markedly increased following sequential stimulation with DC/tumor fusions and anti-CD3/CD28 (28%) as compared with T cell stimulated by anti-CD3/CD28 (11%) or fusions alone (10%) (n = 6) (Fig. 8C). As compared with fusion cells alone, sequential stimulation with DC/breast carcinoma fusions and anti-CD3/CD28 resulted in a 5- and 4-fold increase of CD4+CD25+ T cells that coexpressed CD69 (Fig. 8D) and IFN-γ (Fig. 8E). In contrast, an ~5-fold increase of regulatory T cells was also observed as manifested by an increase in CD4+CD25+ T cells that expressed FOXP3 (Fig. 8F). These results suggest that fusion-mediated stimulation followed by anti-CD3/CD28 expansion induces increased levels of both activated and regulatory T cells.
DC based vaccines represent a promising approach to stimulating antitumor immunity and disease response (2, 49-52). Strategies in which individual tumor Ags are loaded onto DCs are limited by the relative lack of defined immunogenic targets and the risk of immunologic escape through down-regulation of the target Ag. In contrast, DC/breast cancer fusions effectively present a broad array of tumor Ags (12, 16). Endogenously and internalized Ags are presented in the context of the MHC class I and II pathways resulting in a balanced helper and CTL response (53). In a phase I/II trial, 23 patients with metastatic breast and renal carcinoma underwent vaccination with partially mature DCs fused with autologous tumor cells harvested from sites of accessible tissue (17). Fusion cells demonstrated coexpression of tumor-specific Ags, such as MUC1 and DC-derived costimulatory molecules. Vaccination resulted in antitumor immune responses in 10/18 evaluable patients as manifested by an increase in IFN-γ following ex vivo exposure to tumor lysate. Two patients demonstrated disease regression and six patients had stabilization of metastatic disease. Therefore, although the vaccination with DC/breast cancer fusions stimulated antitumor immune responses in a majority of patients, only a subset demonstrated a clinically meaningful disease response.
A major challenge to developing an effective cancer vaccine strategy is overcoming the intrinsic immune deficiencies that limit immunologic response in tumor-bearing patients. Two central elements of tumor-mediated immune suppression include inhibition of DC maturation and the increased presence of regulatory T cells (7-9, 21-23). A potential concern is that tumor cells in the DC/breast carcinoma fusion preparation may inhibit DC development and Ag presentation and induce the expansion of regulatory T cells that would subsequently blunt vaccine response. In previous studies, vaccination with Ag-pulsed immature DCs induced tolerance in Ag-specific T cells (5). In contrast, fusion of immature DCs with multiple myeloma cells resulted in further maturation of the DC fusion partner (54).
In the present study, fusion of DC with breast carcinoma cells resulted in enhanced expression of the costimulatory markers CD80 and CD86, and the maturation marker CD83. Fusion cells generated with immature and mature DCs demonstrated similar levels of maturation, suggesting that the fusion process itself promotes DC activation. Significant expression of IL-12 was observed in both populations consistent with their role as potent APCs with the capacity to stimulate primary immune responses. Expression of CCR7 by the fusion cell populations supports their capacity to migrate to sites of T cell traffic in the draining lymph node. DC/breast carcinoma fusions potently stimulated autologous T cell proliferation with associated secretion of high levels of IFN-γ. Noteworthy, coexpression of IL-12 and IL-10 was observed in the immature and mature DC/breast carcinoma fusion cells. This suggests that the DC/tumor fusions exhibit a mixed phenotype with the capacity to deliver both stimulatory and suppressive signals.
We subsequently examined the effect of DC/breast carcinoma fusions on the relative expansion of activated as compared with regulatory T cells. Stimulation with fusion cells resulted in an increase of CD4+CD25+ cells. Immunophenotyping of this population revealed the presence of activated (CD69+) as well as inhibitory (CTLA-4+, FOXP3) T cells. Stimulation with DC/breast carcinoma fusions resulted in the expansion of CD4+CD25+low and CD4+CD25+high cells, the latter of which demonstrated characteristic findings of regulatory T cells including uniform expression of FOXP3 and suppression of T cell responsiveness. In concert with these findings, a relative increase in both IFN-γ and IL-10 producing cells was observed.
Regulatory T cells play a significant role in mediating tolerance to self Ags in the normal host (55). In patients with malignancy, their increased presence is thought to mediate tumor-associated suppression of host immune responses (18-20). Precise definition of regulatory cells is complex as many markers such as GITR and CD25 are shared between regulatory and activated T cell populations. Regulatory cells are identified by a panel of markers, including CD25high, GITR, CTLA-4, and FOXP3, lack of response to mixed lymphocyte reactions, and the ability to suppress autologous T cell responses in vitro (56-58). DC vaccines may paradoxically increase the presence of regulatory cells that subsequently blunt vaccine response (59). In contrast, depletion of regulatory cells is associated with enhanced vaccine response in diverse tumor models (24, 25). Noteworthy, in a clinical trial, vaccination with DCs pulsed with tumor-specific RNA generated antitumor immunity only when administered in conjunction with an agent (ontak) that eliminates circulating CD25+ cells (25).
Animals models have demonstrated that coadministration of IL-12 and IL-18 promotes T cell activation and augments the efficacy of the DC-based cancer vaccines (14, 32-42). Another strategy to minimize the effect of regulatory T cells is through the activation of innate immunity by ligation of the TLRs (26-31). Administration of CpG ODN to activate TLR9 was shown to overcome the immunosuppression resulting from an expanding tumor burden, decrease the presence of regulatory cells and promote vaccine response. In the present study, addition of the TLR 9 agonists (CpG), IL-12 and IL-18 reduced the levels of regulatory T cells following fusion-mediated stimulation.
In an effort to further enhance vaccine-mediated expansion of tumor reactive lymphocytes, we subsequently examined the effect of sequential stimulation with DC/breast carcinoma fusions and anti-CD3/CD28. CD3/CD28 ligation results in activation of the diverse signaling pathways including those mediated by NFκB (60, 61). Exposure to anti-CD3/CD28 results in T cell expansion and restoration of the complexity of the T cell repertoire in patients with malignancy and HIV (46, 62-64). Adoptive immunotherapy with anti-CD3/CD28 expanded T cells have been examined in patients with renal carcinoma and those undergoing donor lymphocyte infusions after allogeneic hematopoietic stem cell transplantation (65, 66). Therapy was well tolerated and associated with disease response in a subset of patients. However, CD3/CD28 ligation may also deliver an inhibitory signal that promotes the expansion of T cells with an immunosuppressive phenotype. The nature of the T cell response is dependent on the other signals present at the time of stimulation (43). Stimulation with anti-CD3/CD28 alone or in the presence of inhibitory cytokines may induce the expansion of regulatory T cells (67).
We hypothesized that sequential stimulation with DC/breast carcinoma fusions followed by anti-CD3/CD28 would induce an Agspecific primary response with associated inflammatory cytokines that would facilitate CD3/CD28 expansion of the activated T cell compartment. In the present study, this pattern of stimulation resulted in the dramatic expansion of tumor reactive T cell populations far in excess to that seen with either modality alone or when T cells were exposed to anti-CD3/CD28 before the DC/breast carcinoma fusions. The resultant T cell population primarily manifested an activated phenotype.
In summary, DC/tumor fusions exhibit characteristics of potent APCs and stimulate tumor-reactive T cells in vitro. However, fusion cells also induce the expansion of regulatory T cells that potentially inhibit vaccine response. Addition of IL-12, IL-18, and CpG ODN biases the T cell response toward an activated phenotype, limiting the influence of regulatory cells. Ligation of the CD3/CD28 markedly stimulates expansion of tumor-specific memory effector cells. As such, these strategies provide a promising strategy to enhance vaccine efficacy and provide an important platform for adoptive immunotherapy with activated T cells.
1This work was supported by the Department of Defense Grant DAMD17-03-1-0487 and in part by Dana-Farber Cancer Institute/Harvard Cancer Center Ovarian Cancer Specialized Program of Research Excellence Grant P50CA105009-04.
3Abbreviations used in this paper:
The authors have no financial conflict of interest.