It is clear that dendritic cells (DCs) are essential for priming of T cell responses against tumors. However, the distinct roles DC subsets play in regulation of T cell responses in vivo are largely undefined. In this study, we investigated the capacity of OVA-presenting CD4−8−, CD4+8−, or CD4−8+ DCs (OVA-pulsed DC (DCOVA)) in stimulation of OVA-specific T cell responses. Our data show that each DC subset stimulated proliferation of allogeneic and autologous OVA-specific CD4+ and CD8+ T cells in vitro, but that the CD4−8− DCs did so only weakly. Both CD4+8− and CD4−8+ DCOVA induced strong tumor-specific CD4+ Th1 responses and fully protective CD8+ CTL-mediated antitumor immunity, whereas CD4−8− DCOVA, which were less mature and secreted substantial TGF-β upon coculture with TCR-transgenic OT II CD4+ T cells, induced the development of IL-10-secreting CD4+ T regulatory 1 (Tr1) cells. Transfer of these Tr1 cells, but not T cells from cocultures of CD4−8− DCOVA and IL-10−/− OT II CD4+ T cells, into CD4−8+ DCOVA-immunized animals abrogated otherwise inevitable development of antitumor immunity. Taken together, CD4−8− DCs stimulate development of IL-10-secreting CD4+ Tr1 cells that mediated immune suppression, whereas both CD4+8− and CD4−8+ DCs effectively primed animals for protective CD8+ CTL-mediated antitumor immunity.
Dendritic cells (DCs)3 efficiently collect, transport, and present Ags to T cells. Although all DC express multiple surface markers related to their Ag processing and T cell stimulatory functions (e.g., MHC class II, CD40, CD80), they in fact comprise heterogeneous subsets that differ substantially in other surface markers, as well as their developmental origin and physiology. Murine splenic CD11c+ DCs were originally characterized as being either CD8+ or CD8− cells, with both reportedly priming allogenic CD4+ and CD8+ T cells in vitro (1) and Ag-specific CD4+ T cells in vivo (1). Splenic CD8+ DCs reportedly elicit Th1 and CTL responses via an IL-12-dependent mechanism in vivo (2), whereas CD8− DCs stimulate Th0/Th2 responses through their elaboration of IL-10 (1, 3). CD8+ and CD8− DCs from Peyer’s patches similarly induce Th1 and Th2 responses, respectively (4).
More recently, splenic CD8− DCs have been subdivided into CD4− and CD4+ populations (5). Thus, CD11c+ splenic DCs comprise three distinct subsets, with CD4+8− cells representing ∼50% of splenic DCs and CD4−8+and CD4−8− DCs each ∼25% of the total population (6). CD4−8+ DCs have been reported as variably effective (7, 8) stimulators of allogeneic CD8+ T cell responses and CD4−8− and CD4+8− DCs as more effective in stimulating CD4+ T cell responses (8). In vivo, CD4−8+ and CD4−8− DCs can efficiently prime male Ag-specific CTLs, whereas CD4+8− DCs do so only weakly (7). Splenic DCs can also express tolerogenic phenotypes; CD4+8− DCs reportedly mediate tolerance or bystander suppression against diverse T cell specificities (9), while CD4−8+ DCs can induce tolerance to tissue-associated Ags (10). The reasons for the different (i.e., immunogenic vs tolerogenic) results observed using the different DC subsets are at present unclear.
Based on the known critical roles of DCs in induction of primary immune responses, these cells have been used in DC-based cancer vaccines (11). However, given the discrepant immunological functions of the various DC subsets, with their potentials for adverse effects in the context of tumor immunity, we thought it important to critically test the capacity of each DC subset to prime antitumor responses. In this study, we addressed the distinct roles of each of these DC subsets in stimulation of OVA-specific CD4+ and CD8+ T cell responses in vitro and in priming of OVA-specific CTLs as well as antitumor immune responses in vivo.
Materials and Methods
Cell lines, Abs, cytokine, peptides, and animals
The OVA-transfected MO4 cell line was kindly provided by Dr. P. Srivastava (University of Connecticut School of Medicine, Farmington, CT). EL4 T cell lymphoma, EG7 (i.e., OVA-transfected EL4 cells) and LB27 B cell hybridoma expressing I-ab were obtained from the American Type Culture Collection. MO4 and EG7 tumor cells were maintained in DMEM plus 10% FCS and 0.5 mg/ml G418 (Invitrogen Life Technologies). Biotin- and FITC-conjugated monoclonal anti-mouse I-Ab (AF6-120.1), CD4 (GK1.5), CD8 (53-6.7), CD11c (HL3), CD54 (3E2), CD80 (16–10A1), CD86 (GL1), F4/80 (MCAP497), biotin-conjugated anti-CD25 (7D4), CD69 (H1.2F3), Vα2Vβ5+TCR (MR9-4) Abs, and those related FITC-conjugated isotype Abs described above were obtained from BD Pharmingen. FITC- and R-PE-conjugated streptavidin were obtained from Caltag Laboratories. Recombinant mouse GM-CSF and IL-4 were obtained from R&D Systems. The H-2Kb- and I-Ab-specific OVAI (OVA257–264, SIINFEKL) and OVAII (OVA323–339, ISQAVHAAHAEINEAGR) peptides (12) were synthesized by Multiple Peptide Systems. Wild-type, CD4, and IL-10 knockout (KO) C57BL/6, wild-type BALB/c, and the OVA-specific TCR-transgenic OT I and OT II mice (12) were obtained from The Jackson Laboratory.
Homozygous IL-10−/− OT II mice were generated by backcrossing the IL-10−/− mice onto the OT II background for three generations; homozygosity was confirmed using mouse genomic DNA by PCR according to the company’s protocol. All animal experiments were conducted according to the guidelines of the Canadian Council for Animal Care.
Isolation of spleen DCs
This protocol is a modified version of that described by Livingstone and Kuhn (13). Briefly, spleens were injected with DMEM containing 1 mg/ml collagenase (Worthington Biochemical), cut into small fragments, and digested in the above-described enzyme solution for 45–60 min at 37°C. Single-cell suspension was prepared by pressing the digested tissues through a stainless mesh. After the RBC were lysed with Tris-NH4Cl, the spleen cells were washed once in PBS and resuspended in RPMI 1640 plus 10% FCS and 50 μM 2-ME (complete medium) and incubated at 37°C in 100 × 20-mm petri dishes (one spleen equivalent per dish). After 90 min at 37°C, nonadherent cells were removed by gentle washing three times with prewarmed normal saline. Adherent cells were harvested by using trypsin/EDTA (Invitrogen Life Technologies). Spleen DCs were further purified from these adherent cells by incubating them with the biotin-conjugated anti-CD11c Ab and then the anti-biotin Ab-coupled microbeads, and then passing them over a MACS LS column (Miltenyi Biotec) according to the company’s instruction. The microbead-bound cells were fresh CD11c+ spleen DCs. For OVA protein pulsing, adherent cells were cultured overnight in AIM V medium (serum-free lymphocyte medium; Invitrogen Life Technologies) plus GM-CSF (15–20 ng/ml) and OVA protein (0.1 mg/ml) (Sigma-Aldrich). The next day, nonadherent cells were harvested and high-density cells were removed by using Histopaque 1083 (Sigma-Aldrich). The cells were washed twice in 0.5% BSA in PBS and used for isolation of spleen DC subsets.
Purification of spleen DC subsets
DCs subsets were purified as previously described (1) with modification. Briefly, purified overnight cultured nonadherent cells were incubated with the biotin-conjugated anti-CD8 Ab and then with anti-biotin Ab-coupled microbeads, and passed over a MACS LS column (Miltenyi Biotec) according to the company’s instruction. The microbead-bound cells were CD4−8+ spleen DCs. The flow-through cells were then incubated with biotin-conjugated anti-CD4 Ab and then repeated the same procedure as described above for isolation of CD8+ spleen DCs. The bound cells were CD4+8− spleen DCs. The flow-through cells were then passed through a LD column to completely remove any residual cells expressing CD4 and CD8 markers. The final flow-through cells were incubated with the biotin-conjugated anti-CD11c Ab, and the procedure described above was repeated. The bound cells were CD4−8− spleen DCs. The purified CD4+8−, CD4−8+, and CD4−8− spleen DCs were used for the following studies. DCs pulsed with OVA protein were termed as OVA-pulsed DC (DCOVA).
Preparation of DCOVA-activated CD4+ T cells
Naive OVA-specific CD4+ T cells from OT II or OT II/IL-10−/− mice were obtained by passage of splenocytes through nylon wool columns (14). The T cells were fractionated by negative selection using anti-mouse CD8 (Ly2) paramagnetic beads (Dynal Biotech) according to the manufacturer’s protocols to yield populations that were >98% CD4+/Vα2Vβ5+. For activation, the naive CD4+ T cells (2 × 105 cells/ml) were stimulated for 5 days with purified DCOVA subsets (1 × 105 cells/ml) in the presence of IL-2 (10 U/ml) or anti-TGF-β Ab (0.5 μg/ml; R&D Systems) and then isolated from Ficoll-Paque (Amersham Biosciences) density gradients (14).
Analyses of phenotype and cytokine profile
All fresh spleen DCs, purified overnight-cultured spleen DC subsets, and T cells were analyzed by flow cytometry using marker-specific and isotype-matched control Abs as noted (14). The values of mean fluorescence intensity (MFI) for the control and the sample were measured by flow cytometry. The expression index (EI) representing the degree of molecule expression was calculated by dividing the sample MFI with the respective control MFI. For assessment of cytokine production, DC subsets were cultured in the presence of GM-CSF (20 ng/ml) and LPS (1 μg/ml) (15), while DCOVA subset-stimulated CD4+ T cells were restimulated with irradiated (10,000 rad) and OVAII peptide-pulsed LB27 tumor cells (16). One day subsequently, the supernatants were assayed for IFN-γ, IL-4, IL-10, TNF-α, and TGF-β using ELISA kits (R&D Systems) (14).
T cell proliferation assays
Standard T cell proliferation assays were conducted using naive CD4+ or CD8+ T cells purified from BALB/c, OT I, or OT II mice as responder cells and irradiated (4000 rad) DC and DCOVA subsets were used as stimulators, respectively (14).
Splenic lymphocytes (5 × 106) from mice vaccinated with the various DCOVA subsets were cocultured for 4 days in 24-well plates with irradiated (10,000 rad) EG7 cells (2 × 105). The activated T cells were harvested and used as effector cells against radiolabeled EG7 or EL4 target cells in 51Cr release assays (14).
For investigation of antitumor immunity, C57BL/6 mice (n = 8/group) were vaccinated s.c. with 1 × 106 CD4+8−, CD4−8+, or CD4−8− DCOVA and then challenged s.c. 10 days later with 1 × 105 MO4 tumor cells. When investigating the role of CD4+ and CD8+ T cells in antitumor responses, CD4+ and CD8+ T cells were depleted by i.v. injection of 0.5 mg anti-CD4 and anti-CD8 Ab at days −3, 0, and 3 relative to the DC immunization, respectively (14). In all experiments, irrelevant isotype-matched rat Abs were used as controls, and target cell depletion was confirmed by FACS analysis of the circulating T cells. To demonstrate that the anti-CD4 Ab effects (above) were not dependent on a coincidental targeting of CD4+ DCs, in some experiments CD4 KO C57BL/6 mice were used in place of the anti-CD4 Ab-treated mice.
In the studies to confirm that regulatory T (Tr) cells generated in vitro and in vivo were functional in vivo, OT II or OT II/IL-10−/− CD4+ T cells, which had been activated by coculture with freshly purified DCOVA, and CD4+ T cells, which were purified from mice immunized with CD4−8− DCOVA for 7–9 days by using the method described above for purification of CD4+8− DCs with MACS beads, were injected i.v. into mice (2 × 106/mouse) vaccinated with a fully protective dose (i.e., 1 × 106) of CD4−8+ DCOVA 9 days before. OT II and normal CD4+ T cells activated in vitro and in vivo by CD4+8− and CD4−8+ DCOVA were used as controls. These mice were then challenged s.c. with 1 × 105 MO4 tumor cells 1 day later. Animal mortality was monitored daily for up to 10 wk; for humanitarian reasons, all mice with tumors that achieved a size of 1.5 cm in diameter were sacrificed.
Phenotypic characterization of spleen DCs and DC subsets
Fresh spleen DCs were prepared as described in Materials and Methods and then analyzed for their cell surface expression of a panel of immunologically important molecules by flow cytometry. As shown in Fig. 1,A, they mostly express DC marker CD11c, indicating their nature of DC. They displayed a very low expression of MHC class II (I-Ab) (EI, 2.6) and costimulatory molecule CD80 (EI, 1.6) and CD86 (EI, 1.4), which are closely associated with DC maturation, indicating that they are immature DCs. Three subsets of spleen DCs were purified from the overnight-cultured spleen DCs as described in Materials and Methods. As shown in Fig. 1,B, three purified subsets of DC mostly (>90%) expressed CD11c and all expressed high amounts of I-Ab, CD80, and CD86 molecules, indicating that they are mature DCs. There were two distinct peaks of I-Ab, CD54 and CD80 expression, indicating that the differentiation stages of DC subsets are heterogeneous. There was ∼6–7% of the CD3-positive T cell population, but no F4/80-positive macrophages (17), within the above DC subset preparations as examined by flow cytometry (data not shown). Among the three DC subsets, the CD4−8− DC subset had less expression of MHC class II (I-Ab) (EI, 8.3) and costimulatory molecules CD80 (EI, 10.7) and CD86 (EI, 4.0) compared with the other two DC subsets with a 2- to 3-fold higher EI (Fig. 1,B), indicating that it is a relatively less mature form of mature DCs. As shown in Fig. 2, cytokine secretion profiles among DC subsets are significantly different. After stimulation with LPS, both CD4+8− and CD4−8+ DC subsets secreted a higher level of IFN-γ (0.5–1 ng/ml) and TNF-α (0.2 ng/ml) but a very low level of TGF-β. On the contrary, CD4−8− DC subset secreted a much higher level of TGF-β (0.55 ng/ml) (Student’s t test; p < 0.01 vs CD4+8− DCs or CD4−8+ DCs) and a very low level of IFN-γ and TNF-α.
Stimulation of allogeneic T cells in vitro by DC subsets
DCs are potent stimulators of primary MLRs and are able to induce the proliferation of allogeneic T cells in vitro (14). Thus, we compared the ability of three DC subsets in stimulation of primary 3-day MLRs against allogeneic T cells. All three DC subsets demonstrated the ability to stimulate purified CD4+ and CD8+ T cells from BALB/c mice in the 3-day MLRs. Among them, the CD4−8+ DC is the most effective type of DCs in stimulation of proliferation of unfractionated allogeneic T cells, whereas both CD4+8− and CD4−8− DC subsets showed similar stimulation activity (data not shown). When testing on T cell subsets, CD4−8+ DCs are also the most effective in stimulation of both allogeneic CD4+ and CD8+ T cells, whereas CD4+8− DCs are only more effective in stimulation of CD8+ T cells than CD4−8− DCs (Fig. 3, A and B).
Stimulation of tumor Ag-specific T cells in vitro by DC subsets
We next checked and compared their abilities in presentation of tumor Ag OVA instead of alloantigens. For this purpose, we used spleen DCs subsets pulsed with OVA protein for stimulation of transgenic OT II CD4+ and OT I CD8+ T cells in vitro in the 3-day T cell proliferation assay. Accordingly, we found that the results were very similar to those of MLRs. The CD4−8+ DC subset had a higher capacity of stimulating CD4+ and CD8+ T cell proliferation than the other two subsets in the 3-day T cell proliferation assay (Student’s t test; p < 0.05). Again, the CD4−8− DC subset is the weakest inducer of T cell proliferation, probably due to its relatively low maturity. In addition, we also found that the efficiency of T cell proliferation by both CD4+8− and CD4−8− DC subsets peaked at day 3 of stimulation, whereas the efficiency of T cell proliferation by the CD4−8− DC subset peaked at day 5 of stimulation. Interestingly, the CD4−8− DC subset became the strongest inducer in stimulation of both OT II CD4+ and OT I CD8+ T cells in vitro in the 5-day T cell proliferation assay (Fig. 3, C and D).
CD4+8− and CD4−8+ DCs prime CTL-mediated antitumor immune responses in vivo
Next, we wished to test the ability of the various DC subsets for in vivo CTL priming. Splenic lymphocytes from mice immunized with DCOVA subsets were cocultured with irradiated EG7 tumor cells expressing OVA for 4 days and harvested. These T cells are called CTLs. To assess their cytotoxic activities, we conducted a chromium release assay by using these CTLs as effector cells and the 51Cr-labeled EG7 tumor cells as target cells. As shown in Fig. 4 A, CTLs derived from mice immunized with CD4+8− and CD4−8+ DCOVA showed similar cytotoxic activity against EG7 tumor cells. At an E:T ratio of 50, the specific killing activities are all ∼50%, but essentially none of the OVA-negative EL4 cells. In contrast, T cells derived from mice immunized with CD4−8− DCOVA did not show any OVA-specific cytotoxic activity, indicating that the tumor-specific CTL responses in this group of mice were inhibited.
To examine whether DC subsets are capable of inducing antitumor immunity, we vaccinated mice with DCOVA subsets and 10 days later challenged the mice with MO4 tumor cells (1 × 105 cells per immunized mouse). As shown in Fig. 4 B, MO4 tumor cell challenges were invariably lethal within 4 wk after implantation for the control mice vaccinated with PBS, whereas two of eight mice vaccinated with unfractioned splenic DCOVA were protected against MO4 tumor challenge. Most (seven of eight) animals immunized with either CD4+8− or CD4−8+ DCOVA were protected for at least 9 wk, whereas all of the mice immunized with CD4−8− DCOVA died within 5 wk of tumor inoculation.
To study the immune mechanisms in the protective immunity, immunized mice were depleted of their CD4+ T cells using the anti-CD4 Ab before MO4 tumor challenge. As shown in Fig. 5, A and B, the immune protections against MO4 tumor challenge dramatically dropped from 100% to 25% and 0%, respectively, in CD4+ T cell-depleted mice immunized with CD4+8− and CD4−8+ DCOVA. Because the use of anti-CD4 Ab for CD4+ T cell depletion in vivo may also affect and eliminate CD4+8− DCs, we then repeated the CD4+8− DCOVA immunization experiment in CD4 KO mice. As shown in Fig. 5 B, the immune protection became 25% in this group of mice. These results clearly indicate that CD4+ Th cells play an important role in CD4+8− and CD4−8+ DC-primed OVA-specific immune responses. In addition, the immune protection against MO4 tumor challenge dramatically dropped from 100% to 0% in CD8+ T cell-depleted mice immunized with CD4+8− and CD4−8+ DCOVA, indicating that CD8+ CTLs are the major effector cells in the antitumor immunity derived from CD4+8− and CD4−8+ DCs.
CD4−8− DCs prime tolerant immune responses against tumors in vivo
In contrast, CD4−8− DC vaccination did not show any immune protection against the challenge of 0.1 × 106 MO4 tumor cells. All of the mice immunized with CD4−8− DCOVA died within 5 wk after tumor inoculation (Fig. 5 C), confirming that the tumor-specific CTL responses in this group of immunized mice were lacking. To investigate whether CD4+ Tr cells induced by CD4−8− DCs are responsible for the lack of CTL responses, we conducted the above animal studies in CD4+ T cell-depleted mice by using the anti-CD4 Ab. Surprisingly, we found that 88% of mice significantly prolonged their survival (Student’s t test; p < 0.05) and 50% of mice were eventually protected from tumor challenge in CD4+ T cell-depleted mice immunized with CD4−8− DCOVA, compared with the wild-type mice. These results were further confirmed by using CD4 KO mice, which showed that 75% of the immunized mice were protected from tumor challenge. These results clearly indicate that CD4+ Tr1 cells induced by CD4−8− DCs are most likely responsible for the in vivo immune suppression against MO4 tumor cells.
CD4−8− DCs induce CD4+ Tr1 differentiation in vitro
Since DC subsets may be specialized to prime different CD4+ T cell responses (18), we examined the pattern of cytokine secretion of CD4+ T cells activated by OVA protein-pulsed DC subsets by ELISA. We first confirmed that these activated T cells were CD4 positive and displayed the clonotypic Vα2Vβ5 OVA TCR, as well as the T cell activation markers CD25 and CD69 (data not shown). We then examined their cytokine profiles. We found that the activated CD4+ T cells from the CD4+8− or CD4−8+ DCOVA cocultures secreted similarly high levels of IFN-γ, but relatively little or no IL-4 or IL-10 (Fig. 6 A), indicating a Th1 phenotype. However, the CD4+ T cells activated by TGF-β-secreting CD4−8− DCOVA secreted high levels of IL-10 (1.75 ng/ml) and substantial amounts of IFN-γ (2.1 ng/ml), indicating a Tr1 phenotype.
CD4+ Tr1 cells inhibit the antitumor immunity through IL-10
Because Tr1 cells suppress immune responses (19), we next assessed whether they also inhibit the antitumor immunity in our animal model. We injected these Tr1 cells into CD4−8+ DCOVA-immunized mice before tumor challenge. As shown in Fig. 6,B, CD4−8− DCOVA-induced OT II CD4+ Tr1 cells completely inhibited the immune protection of CD4−8+ DCOVA immunization, whereas CD4+8− or CD4−8+ DCOVA-activated Th1 cells showed enhanced protection against MO4 tumor challenge when compared with the CD4−8+ DCOVA control. To confirm the above findings, we also isolated the CD4+ T cells from CD4−8− DCOVA-immunized immunized mice and repeated the same experiment as that for in vitro-activated Tr1 cells. As we expected, we found almost identical results to those for in vitro-cultured Tr1 cells (data not shown), confirming that CD4+ Tr1 cells induced by CD4−8− DCOVA are most likely responsible for the in vivo immune suppression against MO4 tumor cells. To assess whether IL-10 was involved in the inhibition of CD4−8+ DCOVA-induced antitumor immunity, OT II CD4+ T cells (IL-10−/−) were used to coculture with the CD4−8− DCOVA. After 5 days of culture, the Tr1 (IL-10−/−) cells were injected into CD4−8+ DCOVA-immunized mice. As shown in Fig. 6 B, most of the immunized mice were protected against the MO4 challenge and no inhibition was observed, while the normal Tr1 cells completely inhibited the antitumor immunity of CD4−8+ DCOVA.
TGF-β is partially responsible for the induction of Tr1 by CD4−8− DCOVA
It has recently been reported that LPS-stimulated B cells expressing TGF-β induced T cell anergy via activation of Tr cells (18). To explore the mechanism of Tr1 induction, we added anti-TGF-β Ab to the coculture of CD4−8− DCOVA and OT II CD4+ T cells and then tested the cytokine profile of activated CD4 T cells. Interestingly, their cytokine profile became similar to that of CD4−8+ DCOVA-activated Th1 (Fig. 6,A). Their IL-10 secretion was decreased nearly 10 times when compared with their controls. Furthermore, these T cells were used in in vivo study as described above. No inhibition, but a little enhancement of antitumor immune response, was observed in the CD4−8+ DCOVA-immunized mice (Fig. 6 B), indicating that TGF-β was involved in promoting the formation of Tr1 cells.
Because of the critical roles DCs have in induction of primary immune responses, they have been extensively used for DC-based cancer vaccines. It has been shown that DCs, when pulsed with tumor-derived MHC class I-restricted peptides and tumor Ags, are able to induce significant CTL-dependent antitumor immune responses (11). Because the distinct immunological functions of DC subsets may have detrimental effects on the host’s immune responses, it is important to assess the capacity of these DC subsets in priming antitumor immunity.
Splenic DCs have been classified into three subsets (CD8+4−, CD8−4+, and CD4−8− DCs). Although the phenotypic and functional differences between CD8+ and CD8− DC subsets have been extensively studied (1, 2, 3, 4), the differences between CD4+ and CD4− populations within the CD8− DC subset have been less investigated. In this study, we conducted a systemic study on the phenotypic characteristics and the functional differences in stimulation of T cells among the three DC subsets. For the first time, our data showed that CD4−8− DCs secreted marked levels of TGF-β and represent a characteristic consistent with a tolerogenic phenotype (20). In contrast, the CD4+8− and CD4−8+ DCs secreted moderate and high levels of IFN-γ relative to the CD4−8− DCs, respectively, but little or no TGF-β. We then investigated the capacity of three DC subsets pulsed with OVA protein in priming OVA-specific antitumor immune responses in vivo. Our data showed that only CD4+8− DCOVA and CD4−8+ DCOVA could induce the antitumor response and protect the immunized mice from tumor challenge, whereas CD4−8− DCOVA could not stimulate any protective immune response against tumor, indicating distinct in vivo antitumor immune responses derived from vaccination of three DC subsets.
There is now compelling evidence that CD4+ T cells, specialized in suppressing immune responses, play a critical role in immune regulation. Three major populations of Tr cells have been identified based on their distinct phenotype (CD4+25+) or cytokine profile (Tr1 and Th3). The CD4+25+ Tr subset mediates immune suppression in a non-Ag-specific manner, whereas the latter Tr1 and Th3 mediate immune inhibition via production of IL-10 and TGF-β, respectively, but both in an Ag-specific way (21). In this study, the cytokine profiles of CD4+ T cells activated by DCOVA subsets differed markedly. CD4+8− or CD4−8+ DCOVA induce Th1 phenotype response, whereas T cells activated by TGF-β-secreting CD4−8− DCs secreted high levels of IL-10 and substantial amounts of IFN-γ, a characteristic of Tr1 cells (21). Tr1 cells can also be induced by addition of exogenous IL-10 to primary murine T cell cultures or by coculturing T cells with TGF-β/IL-10-expressing “tolerogenic” DCs (19, 22, 23) or by immature DCs (24). In addition, others have also shown that the IL-10-stimulated CD11clow CD45RBhigh tolerogenic DCs, which have a phenotype similar to our CD4−8− DCs, also induced Tr1 cell differentiation and immune tolerance in vivo (25). LPS-stimulated B cells expressing membrane-bound TGF-β have recently been shown to induce T cell anergy (20) via the activation of Tr cells (26). Interestingly, in this study, the addition of anti-TGF-β Ab to CD4−8− DCOVA coculture with OT II T cells converted the cytokine profile of these CD4+ T cells from the Tr1 phenotype to the Th1 phenotype, indicating the critical role of TGF-β of CD4−8− DCs in stimulation of the Tr1 response.
These Tr1 cells also distinct from Th1 or Th2 cells in that they produce high levels of IL-10 and no IL-4, and proliferate poorly upon TCR ligation, suppress immune responses in vitro and in vivo through secreted IL-10 (27). In this study, we also characterized the functional effect of these Tr1 cells in induction of immune suppression in vivo. The CD4+ Tr1 cells from CD4−8− DCOVA/OT II cocultures or purified from CD4−8− DCOVA-immunized mice were transferred into CD4−8+ DCOVA-immunized mice and abolished the CD4−8+ DCOVA-induced antitumor immunity. Unlike the wild-type Tr1 cells (above), the IL-10−/− CD4 OT II T cells activated with CD4−8− DCOVA had little impact on CD4−8+ DCOVA-driven antitumor immunity, clearly implicating the IL-10-producing Tr1 cells as central to the tolerance observed in this model system, and this is consistent with previous reports (26, 27). As expected, when we transferred the control Th1 cells from CD4+8− or CD4−8+ DC/OT II CD4+ T cell cocultures or T cells from coculture with CD4−8− DCOVA in the presence of anti-TGF-β Ab into CD4−8+ DC-vaccinated animals, we observed augmented tumor protection, confirming the critical role of TGF-β of CD4−8− DCs in stimulation of the Tr1 response.
CD4+ Th and Tr cells play an important role in modulation of immune responses by enhancement and suppression of CD8+ CTL responses, respectively (28). In this study, we showed that depletion of CD4+ Th and Tr cells significantly reduced and enhanced antitumor immune responses in CD4+8− or CD4−8+ DC- and CD4−8− DC-immunized mice, respectively. Surprisingly, the immune protection in CD4+ T cell-depleted CD4−8− DC-immunized mice is much stronger than that in CD4+ T cell-depleted CD4+8− or CD4−8+ DC-immunized mice. The immune mechanism behind this phenomenon is currently unknown. The enhanced immunity seen in CD4+ T cell-depleted CD4−8− DC-immunized mice may be partially derived from the capacity of CD4−8− DCs making more IL-12p70 than CD4+8− DCs when stimulated appropriately (29), which is associated with stimulation of CTL responses (30). It has been reported that different spleen DC subsets when pulsed with MHC class I-restricted viral peptide can induce antiviral immunity mediated by Th-independent CTL responses (31). In this study, we also found that our three DC subsets when pulsed with OVAI peptide can all induce protective immunity against MO4 tumor cells, among which the CD4−8− DC subset stimulates the strongest immunity (data not shown), also supporting the above finding.
Discrepant results regarding immune priming vs tolerance have previously been reported for CD4+8− (8, 9) and CD4−8+ DCs (1, 2, 10). In addition, our results of tolerogenic CD4−8− DCs in antitumor immunity are also in contrast to a previous report wherein CD4−8− DCs efficiently stimulated H-Y Ag-specific CTL responses (7). It has been reported recently that the environmental conditions or stimuli under which DCs stimulate T cells critically affect the type of immune responses that ensue (14, 32, 33). This suggests that the discrepancies between our results and those observed in other systems (e.g., autoimmunity, allotransplantation, or antitumor immunity) would likely be attributable to the varying antigenic and environmental conditions in each model system, where different TLRs are stimulated (34).
Taken together, our data unequivocally confirmed that the different DC subsets can function either as stimulators or inhibitors of immune responses. CD4−8− DC stimulated IL-10-secreting CD4+ Tr1-mediated immune suppression, whereas both CD4+8− and CD4−8+ DC induced CD8+ CTL-mediated antitumor immunity. This information has very substantial implications for DC-based approaches to the design of cancer vaccines.
We thank Mark Boyd for flow cytometric analysis.
The authors have no financial conflict of interest.
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
This work was supported by a research grant (MOP 63259) from the Canadian Institute of Health Research, a Bridge Research Fund of Saskatchewan Health Research Foundation, and the Hazel Constance Brooker Research Trust Fund (to J.X.).
Abbreviations used in this paper: KO, knockout; DCOVA, OVA-pulsed DC; Tr, regulatory T; KO, knockout; MFI, mean fluorescence intensity; EI, expression index.