Tumor growth is associated with the inhibition of host antitumor immune responses that can impose serious obstacles to cancer immunotherapy. To define the potential contribution of Qa-1–restricted CD8 regulatory T cells (Treg) to the development of tumor immunity, we studied B6.Qa-1 D227K mice that harbor a point mutation in the MHC class Ib molecule Qa-1 that impairs CD8 Treg suppressive activity. Here, we report that the growth of B16 melanoma is substantially delayed in these Qa-1–mutant mice after therapeutic immunization with B16 melanoma cells engineered to express granulocyte macrophage colony-stimulating factor compared with Qa-1 B6-WT controls. Reduced tumor growth is associated with enhanced expansion of follicular T helper cells, germinal center B cells, and high titers of antitumor autoantibodies, which provoke robust antitumor immune responses in concert with tumor-specific cytolytic T cells. Analysis of tumor-infiltrating T cells revealed that the Qa-1 DK mutation was associated with an increase in the ratio of CD8+ T effectors compared with CD8 Tregs. These data suggest that the CD8+ T effector–Treg ratio may provide a useful prognostic index for cancer development and raise the possibility that depletion or inactivation of CD8 Tregs represents a potentially effective strategy to enhance antitumor immunity. Cancer Immunol Res; 2(3); 207–16. ©2013 AACR.
Recent advances in cancer immunology have generated considerable interest in approaches that interrupt inhibitory pathways that can constrain antitumor immunity (1, 2). For example, definition and analysis of CTLA-4 expression by regulatory and other T cells led to the generation of a CTLA-4 monoclonal antibody (ipilimumab) that substantially extends survival in patients with metastatic melanoma (3, 4) and shows promise for achieving durable remissions of other types of cancer. Although ipilimumab has been approved as a standard-of-care therapy for advanced melanoma since 2011, a relatively small fraction (approximately 1/5) of patients treated with this drug achieved long-term clinical responses, suggesting that targeting additional inhibitory pathways might increase the efficacy of this general therapeutic strategy.
Most current approaches directed at regulatory T cells (Treg) have targeted cells within the CD4 lineage. However, there is increasing evidence that CD8 T cells also include a regulatory cell lineage that inhibits T-cell responses (5). These regulatory CD8 T cells (CD8 Treg) eliminate activated T follicular helper (TFH) cells through recognition of the MHC class Ib molecule, Qa-1 (HLA-E in human) expressed at their surface. This regulatory subset represents about 5% of CD8 T cells and expresses a triad of surface receptors, CD44, CD122, and Ly49, that can be used to purify them (6, 7). Qa-1 knock-in mice (B6.Qa-1 D227K; B6-DK) have been generated to express a point mutation (position 227, D → K) that disrupts binding of Qa-1 to the T-cell receptor (TCR)–CD8 complex of CD8 Tregs (8). These mutant mice develop increased germinal centers, high titers of autoantibodies, and a lethal systemic lupus erythematosus (SLE)-like autoimmune disease (6), highlighting the contribution of this Qa-1–restricted inhibitory interaction to regulation of the immune response and self-tolerance.
These considerations led us to examine the potential contribution of Qa-1–restricted inhibitory pathways to antitumor immunity. Analysis of mice that had been inoculated with granulocyte macrophage colony-stimulating factor (GM-CSF)–expressing B16 melanoma cells revealed robust upregulation of Qa-1 on lymphocytes and tumor infiltration of CD8 Tregs. We reasoned that increased targeting of Qa-1+ TFH cells by CD8 Tregs might inhibit tumor immunity, and disruption of this inhibitory interaction might enhance the protective immune response. We therefore analyzed the response of B6-DK mice challenged with B16 melanoma after vaccination with irradiated GM-CSF–transduced tumor cells. We find that genetic disruption of CD8 Treg activity results in enhanced antitumor immunity that is associated with a robust antibody response to tumor-associated antigens (TAA) that cooperates with CD8 effector T cells (Teff) to constrain tumor growth.
Materials and Methods
C57BL/6J (B6; Jackson Laboratory), B6.Rag2−/− (Taconic), and B6-DK mice [N11; 99% congenic according to single-nucleotide polymorphism (SNP) analysis], OT-I [C57BL/6-Tg(TcraTcrb)1100 Mjb/J], and OT-II [B6.Cg-Tg(TcraTcrb)425 Cbn/J] mice were housed in pathogen-free conditions. All experiments were performed in compliance with federal laws and institutional guidelines, as approved by the Institutional Animal Care and Use Committee (IACUC) of the Dana-Faber Cancer Institute (Boston, MA).
Cell lines and tissue culture
B16F10 (American Type Culture Collection) and GVAX (B16-GM-CSF) were cultured in complete Dulbecco's Modified Eagle Medium (DMEM) with 10% fetal calf serum (FCS). B16-OVA was cultured in complete DMEM with 10% FCS supplemented with 250 μg/mL G418 (Invitrogen). All lines were maintained at 37°C and 5% CO2. All cell lines used were checked for mycoplasma. B16-OVA was authenticated by PCR for OVA expression. No additional authentication was performed.
Reagents and flow cytometry
Single-cell suspensions were stained with target antibodies for 30 minutes in the dark at 4°C in ice-cold fluorescence-activated cell sorting (FACS) buffer (2% FCS, 0.1% NaN3 in PBS), washed, and analyzed using a FACSCalibur or LSR Fortessa (BD Biosciences) and FlowJo software (TriStar). Tumors were digested with collagenase/dispase for 2 hours at 37°C with agitation and fractionated on Nyco-Prep 1.007 separation medium (PROGEN) with centrifugation at 2,000 rpm at room temperature. Lymphoid fraction was collected, washed extensively with ice-cold FACS buffer, and analyzed. Tumor infiltrates were defined as CD45+ cells. Anti-B220, anti-CD44, anti-CD45, anti-CD122, anti-Ly49 C/I/F/H, anti-NK1.1, anti-CD11b, anti-CD25, anti-CXCR5, anti-ICOS, IgG1 isotype, anti–Qa-1b, anti-CD69, anti-Fas, and anti-IgM (BD Bioscience) or anti-CD4, anti-CD8, anti-CD3, anti-CD200, anti-BTLA4, anti–PD-1, anti-FoxP3 (eBioscience), and CD1d tetramer (NIH Tetramer Core Facility at Emory University) were used for cell analysis.
Tumor challenge and treatment
Six- to 12-week-old female B6-WT or B6-DK mice were challenged subcutaneously with 2–5 × 105 B16 (or B16-OVA) tumor cells and immunized with irradiated 1 × 105 GVAX (B16 retrovirally transduced with GM-CSF) subcutaneously on the opposite flank on day 0. Mice were treated by immunization with 5 × 105 irradiated (150 Gy) B16 (or B16-OVA) and 5 × 105 GVAX s.c. on days 7 and 14 after challenge on alternating sides. Tumors were measured 2 to 3 times per week using calipers and tumor volume was presented as (x × y × z)/2 mm3. Mice exhibiting signs of distress or with tumors reaching 2 cm on the longest axis were sacrificed according to IACUC guidelines. Survival plot was created using Prism 6.0 (GraphPad Software, Inc.). In some experiments, mice were vaccinated with irradiated 5 × 105 B16-OVA and 5 × 105 GVAX s.c. 1 week before challenge with 1 × 106 B16-OVA and tumor incidence was recorded. In serum transfer experiments, Rag2−/− hosts were challenged with 1 × 106 B16-OVA and treated with 400 μL serum intraperitoneally on days 0, 8, and 15 after challenge. Mice also received 2 × 106 naïve OT-I Tg CD8 T cells intravenously on day 2 and 1 μg IL15–IL15Rα complex intraperitoneally on days 2, 7, and 14 after challenge. Serum was prepared from either B6-WT or B6-DK mice vaccinated on days 0, 7, 10, and 17 with irradiated 1 × 106 B16-OVA and 5 × 105 GVAX. Serum was collected beginning on day 13 after the first immunization, and equal volumes from each collection date were pooled before treatment. Serum was prepared as described (9), total IgG was obtained by precipitation with ammonium sulfate (Sigma), diluted in PBS, and desalted on 50,000 MW Amicon filters (EMD Millipore).
Detection of NP-specific antibodies by ELISA was performed as described (6). Serum harvested 14 days after immunization with NP19-KLH in complete Freund's adjuvant (CFA) and reimmunization with NP19-KLH in IFA was used as a standard. A 1:1,000 dilution of this immune serum was defined as 100 U/mL. OVA-specific antibodies were detected on ELISA plates coated with 10 μg/mL OVA (Pierce) overnight at 4°C. Total IgG1 was detected by incubating plates with biotinylated anti-mouse IgG1 conjugated to streptavidin peroxidase. Anti-OVA IgG1 standard was from Abcam. Anti–double-stranded DNA (dsDNA) and anti-nuclear antigen (ANA) antibodies in mouse sera were determined by ELISA according to the manufacturer's protocol (Alpha Diagnostic International). Anti-macrophage migration inhibitory factor (MIF) and anti-angiopoietin-2 (Ang-2) IgG antibodies were measured as described (10). Recombinant mouse MIF and Ang-2 were from R&D Systems. IFN-γ in mouse sera was measured using an OptEIA ELISA Kit (BD Biosciences); interleukin (IL)-21 was measured using an ELISA Duo Set (R&D Systems).
Sections from frozen spleens and tumors, 7 μm and 10 μm, respectively, were fixed in cold acetone, air-dried, blocked with 3% bovine serum albumin in PBS and 1 μg/mL Fc block (BD Biosciences) for 30 minutes at room temperature, and stained with antibodies overnight in blocking buffer in the dark at 4°C using a moist chamber. Images were acquired on a Nikon inverted wide-field fluorescence microscope with 100× magnification. Germinal centers in spleen were defined by labeling with phycoerythrin (PE)-conjugated anti-B220 antibodies (BD, RA3-6B2) and fluorescein isothiocyanate (FITC)–conjugated anti–GL-7 (BD, clone GL7) antibodies. IgG deposition in tumor was detected with FITC-conjugated anti-mouse IgG antibody (Sigma), complement membrane-activated complex (MAC) with biotinylated C5b-9 antibody (Bioss), macrophages with F4/80 antibody (eBioscience), desmin with anti-desmin Ab (Abcam), and fibroblast with anti-ERTR7 Ab (Santa Cruz Biotechnology). Data were quantified using ImageJ software and presented as fluorescent area (pixels2) per field of view (FOV) or using NIS Elements software (Nikon Instruments Inc.) and presented as fluorescent area (pixels2) per region of interest (ROI). Vasculature was visualized using PE-conjugated anti-CD31 antibody (eBioscience) and vessel diameter was measured on the longest axis by NIS Elements software.
Adoptive transfer experiments
Naïve CD4+CD25− cells were purified from spleens of OT-II Tg WT or B6-DK mice using a CD4 Cell Enrichment Set and biotinylated anti-CD25 antibody (BD Biosciences). CD4 cells were in vitro polarized to TFH cells with 50 ng/mL IL-21, 20 ng/mL IL-6, 10 μg/mL anti–IL-4, 10 μg/mL anti–IFN-γ, 20 μg/mL anti–TGF-β, and cultured for 5 days in the presence of 1 μg/mL OT-II peptide and antigen-presenting cells. Naïve B cells were isolated from the spleens of B6-WT mice using a B Lymphocyte Enrichment Set (BD Biosciences). B-cell and CD4 cell purity was >95%, according to FACS analysis. CD8 Tregs were isolated from the spleens of B6-WT mice immunized intraperitoneally with 100 μg of keyhole limpet hemocyanin (KLH) in CFA, and reimmunized 7 days later with 100 μg of KLH in incomplete Freund's adjuvant (IFA). CD8 Tregs were first enriched with a CD8 Cell Enrichment Set (BD Bioscience) followed by sorting for CD3+CD8+CD44+CD122+Ly49+ T cells. Rag2−/− mice were adoptively transferred with 1.5 × 106 OT-II TFH cells, 3.5 × 106 naïve B cells, and 1 × 105 CD8 Tregs. Immediately after transfer, mice were immunized intraperitoneally with 100 μg of NP13-OVA in CFA, and after 10 days, mice were reimmunized intraperitoneally with 50 μg of NP13-OVA in IFA and challenged with 1 × 106 B16-OVA s.c. 4 days later. Tumor size was monitored as mentioned above.
Data are presented as mean ± SEM. Statistical analyses were performed using Prism 6.0 software, the Wilcoxon–Mann–Whitney rank sum test for comparison of two groups or conditions, unless otherwise noted. P < 0.05 was considered to be statistically significant (*, P < 0.05; **, P < 0.01; ***, P < 0.001).
Genetic disruption of CD8 Treg activity is associated with reduced melanoma growth and enhanced TFH cell responses
We utilized the B16 melanoma model to investigate the contribution of CD8 Tregs to antitumor immunity (11). Tumor-bearing B6 mice were vaccinated with irradiated B16 melanoma cells engineered to express GM-CSF (GVAX) to induce an immune response to the tumor (12). Following GVAX, we noted substantial upregulation of Qa-1 expression by splenocytes and by tumor-infiltrating lymphocytes (TIL), but not by tumor cells (Fig. 1A). As B16 tumor cells do not express detectable Qa-1, host cells in the spleen and tumor infiltrates represent potential targets of Qa-1–restricted CD8 Tregs. Upregulation of Qa-1 expression by immune cells was associated with the infiltration of B16 tumors by cells expressing markers of the CD8 Treg phenotype (Fig. 1B). Increased proportions of CD8 Tregs within tumor-infiltrating CD8 T cells correlated with rapid tumor growth (∼day 20; Fig. 1B, right). We directly tested the contribution of Qa-1–restricted CD8 Tregs to tumor rejection using Qa-1 knock-in mice (B6-DK) that harbor defective Qa-1–restricted CD8 Treg activity secondary to a Qa-1 point mutation that disrupts TCR recognition of Qa-1–peptide ligands (8). We inoculated B6-WT and B6-DK mice with B16 tumor cells and immunized them with irradiated GM-CSF–transduced B16 cells on day 0, 7, and 14. B6-DK mice showed significantly extended survival and reduced tumor growth compared with B6-WT mice (Fig. 1C).
Further analysis of B6-DK mice revealed increased numbers of TFH cells (CD4+ICOS+CD200+) compared with B6-WT mice (Fig. 1D), consistent with previous findings that immunization of B6-DK mice with foreign antigens results in increased expansion of TFH cells and high titers of autoantibodies (6). Moreover, tumor-infiltrating CD4 T cells in Qa-1–mutant mice displayed a more activated phenotype, as judged by levels of ICOS expression (Fig. 1D; ref. 3). No significant difference was noted in the numbers of CD4 Tregs or NK cells (Fig. 1E). Interestingly, increased intratumoral expansion of effector CD8 T cells and reduced numbers of CD8 Tregs were detected in B6-DK mice compared with B6-WT controls (Fig. 1E). This resulted in a substantial increase in the intratumoral CD8+ Teff–Treg ratio that was associated with the Qa-1 DK mutation.
Vaccination of B6-DK mice results in enhanced antibody responses to TAA
In view of the increased numbers of TFH cells (e.g., Fig. 1D) and germinal center B cells (see below) in tumor-bearing B6-DK mice, we asked whether these Qa-1–mutant mice developed antibody responses to surrogate TAA. We immunized B6-WT and B6-DK mice with irradiated GM-CSF–transduced B16 tumor cells that expressed an ovalbumin transgene (B16-OVA; ref. 13). Two to four weeks later, we detected high titers of OVA-specific antibody in the sera from B6-DK, but not B6-WT mice (Fig. 2A). Increased anti-TAA antibody formation was associated with enhanced germinal center formation in B6-DK mice (Fig. 2B). Vaccination of B6-DK mice with irradiated B16-OVA and GVAX 7 days before inoculation with B16-OVA tumor cells also induced high titers of anti-OVA antibodies. This antibody response was maintained during the course of B16 tumor growth and correlated with increased tumor protection (Supplementary Fig. S1), suggesting that antibodies induced by vaccination might have durable antitumor activity. Moreover, higher titers of anti-OVA antibodies in B6-DK mice were also detected when mice were first injected with B16-OVA tumor cells 7 days before immunization with irradiated B16-OVA and GVAX, supporting the idea that the enhanced response to TAA in B6-DK mice might have a therapeutic value (Fig. 2C).
To evaluate the contribution of TFH cells to antitumor antibody responses in Qa-1–mutant mice, we adoptively transferred in vitro polarized TFH cells from B6-WT OT-II or B6-DK OT-II TCR Tg mice into Rag2−/− hosts along with naïve B6-WT B cells and CD8 Tregs. Following inoculation with B16-OVA cells and challenge with NP-OVA, recipients of TFH cells from B6-DK mice produced substantially higher titers of both total anti-NP (anti-NP23 IgG1) and high affinity (anti-NP4 IgG1) antibodies and anti-OVA antibodies than recipients of TFH cells from B6-WT donors, and this response was associated with reduced tumor growth (Fig. 3A and B). We again observed enhanced expansion of TFH cells (but not CD8 Tregs) in recipients of B6-DK TFH cells (Fig. 3C). The expansion of TFH cells was accompanied by increased levels of IL-21, but not IFN-γ in the sera of B6-DK mice (Fig. 3D), indicating that the expanded T cells were producing a canonical TFH cytokine.
Next, we analyzed the repertoire of antibodies present in the serum of GVAX-vaccinated B6-WT and B6-DK mice. We found increased titers of autoantibodies in the sera of B6-DK mice, including anti-dsDNA antibodies (Fig. 4A). The same trend was observed after treatment of tumor-bearing mice with GVAX (Fig. 4B). In addition, we noted several types of autoantibodies that have been reported to have antiangiogenic properties (10, 14, 15) in a portion of the B6-DK mice, but not B6-WT mice (Fig. 4C). These antibodies were induced upon GVAX vaccination and were not detectable in naïve B6-WT or B6-DK mice. To determine whether the tumor vasculature was affected by the therapeutic antibodies in B6-DK mice, we performed histology on tumor sections and immunostaining with CD31 antibodies. Although the number of vessels was similar, the tumor vessels appeared smaller in tumors of B6-DK than in B6-WT controls (Fig. 4D), suggesting that increased vasculature pathology was associated with increased antibody responses of B6-DK mice, which may contribute to delayed tumor growth.
Most analyses of protective tumor immunity have focused on cell-mediated immunity, including the host cytotoxic response. Our analysis of enhanced antitumor immunity in animals with defective CD8 Treg activity suggested a role for autoantibody responses, including antibodies to TAA. To determine whether antibodies produced after GVAX immunization of B6-DK mice contributed to antitumor immunity, Ig was prepared from the serum of B6-WT or B6-DK mice after GVAX immunization. Rag2−/− mice were inoculated with B16-OVA and treated with 400 μL of serum-derived Ig on days 0, 8, and 15. Mice also received 2 × 106 naïve OT-I cells on day 2 and 1 μg IL15/IL15Rα complex on days 2, 7, and 14 after tumor inoculation as a source of activated CD8 T cells. We found that 60% of the mice that received antibodies from B6-DK mice remained tumor free, whereas all mice that received serum Ig from WT controls developed tumors (Fig. 5A). This antitumor response did not depend on the contribution of anti-OVA antibody alone because the provision of high titers of anti-OVA Ig (500 μg anti-OVA IgG1) to Rag2−/− mice without T cells had no detectable effect on tumor growth (Supplementary Fig. S2). Further analysis of B16 tumors revealed increased IgG deposition in the tumors of B6-DK mice after treatment with GVAX (Fig. 5B), which colocalized with the C5b-C9 complement MAC (Fig. 5B) and was associated with macrophage infiltrates (Fig. 5C); Ig did not colocalize with other tumor stromal components, including CD31+ and ERTR-7+ cells (Fig. 5D). We also used a polyclonal antibody to the desmin stromal protein that cross-reacts with single- and double-stranded DNA (16) for additional histologic analysis (Supplementary Fig. S3). Immunofluorescence analysis using anti-desmin antibody showed colocalization with IgG deposits in addition to the presence of filamentous structures normally detected by this antibody (Fig. 5D). Areas containing colocalized anti-desmin antibody accounted for a substantial fraction of the intratumoral IgG foci (∼30%–50%), suggesting that a significant proportion of intratumoral Ig may represent anti-DNA antibodies derived from sera of tumor-bearing B6-DK mice.
We report here that antitumor immunity against melanoma is enhanced by the genetic disruption of CD8 Treg activity that is associated with the expansion of TFH cells and germinal center B cells, the generation of anti-TAA antibodies, and delayed tumor growth. These findings concerning the antitumor response are consistent with previous findings that targeting of TFH cells by CD8 Treg constrains the expansion of TFH cells and the development of high-affinity antibody and autoantibody responses. Excessive production of IL-21 by activated TFH cells in B6-DK mice may also stimulate a CD8 T-cell response to exert antitumor activity (17).
The therapeutic activity of the antibody response mounted by B6-DK mice was confirmed from studies of tumor-bearing hosts infused with IgG from immunized tumor-bearing B6-DK, but not from B6-WT mice. Transfer of IgG from the former, but not the latter donors resulted in significant protection from lethal tumor growth in adoptive hosts (Fig. 5A). The antibody response in B6-DK mice included anti-dsDNA, anti-MIF, and anti–Ang-2 specificities after GVAX treatment and tumor challenge (Fig. 4). Increased IgG deposition in tumors of B6-DK mice was associated with tumor tissue destruction and often colocalized with MACs (C5b-C9) that have the potential to lyse tumor cells. Whether antibodies generated in this setting directly target B16 melanoma or act through the activation of tumor-infiltrating immune cells, such as macrophages, is not clear. NK-mediated cytolysis of B16 melanoma does not appear to be involved in this process because NK cells did not colocalize with intratumoral IgG and we did not observe antibody-dependent cell-mediated cytotoxicity against B16 cells after incubation with B6-DK–derived antibody.
Considerable attention has been given to the association of inflammatory and autoimmune responses and the development of cancers (18–22). However, the influence of chronic autoimmunity on the natural history of solid tumors is not well understood. For example, although patients with SLE have decreased risk of breast cancer and melanoma (23), the underlying mechanisms and biologic significance of these observations are unclear. It may be relevant that longitudinal analyses of sera from long-term surviving melanoma patients after vaccination with GM-CSF–secreting autologous melanoma cells have revealed that the generation of autoantibodies was associated with increased tumor destruction (15).
These findings raise the intriguing possibility that interruption of CD8 Treg activity may act in part through enhanced antibody responses to “classical” autoimmune target antigens, including dsDNA. While the full repertoire of antibodies generated in B6-DK mice upon GVAX treatment and their relative contribution to antitumor immunity has not been determined, the production of high levels of anti-dsDNA antibody in tumor-bearing B6-DK mice after GVAX treatment was associated with protective antitumor immunity. Although the development of anti-dsDNA antibody is considered a pathogenic sign in the context of SLE, the appearance of these autoantibodies in patients with cancer has been associated with improved clinical outcome (24). Anti-dsDNA antibodies may affect tumor growth through direct binding and induction of apoptosis (25, 26). Cell-penetrating anti-DNA antibodies, for example, inhibit DNA single- and double-stranded DNA repair in tumor cells and sensitize them to DNA-damaging therapy (27). Anti-dsDNA antibody complexed to dsDNA can also stimulate B cells and dendritic cells in a TLR9-dependent manner to produce increased complement-fixing antibodies and IFN-α, respectively, which can have inhibitory effects on malignancy (28, 29). These considerations support the notion that generation of a broad-based range of autoantibodies, including dsDNA, in the absence of CD8 Treg activity may contribute to the enhanced anti-melanoma responses noted here as well as to the response against a wide range of tumors.
The ability of antibodies specific for intracellular rather than cell surface molecules to mediate antitumor immunity is not well understood. The pathogenic effects of antibodies to the cytoplasmic enzyme glucose-6-phosphate isomerase (GPI) in the K/BXN arthritis mouse model may be instructive. Here, anti-GPI antibodies initiate local inflammatory changes leading to increased extracellular GPI and enhanced vascular permeability to serum autoantibodies and inflammatory cells (30, 31). In the present study, antibody-mediated antitumor effects depended on the cooperative action of tumor-specific CD8 T cells (Fig. 5A). Induction of inflammatory foci in the tumor environment may enhance the release of intracellular antigens (e.g., dsDNA), resulting in increased permeability of the vasculature and mobilization of antigen-specific CD8+ cytolytic T cells into the tumor sites. This coordinated humoral and cellular response may be essential for optimal antitumor activity. Indeed, a combined antibody and CD8+ antitumor immune response to the intracellular melanoma antigen NY-ESO-1 has been correlated with clinical benefit in patients with advanced melanoma treated with anti-CTLA-4 antibody (ipilimumab; ref. 32). Increased ICOS+ CD4 cells within the tumor and enhanced IL-21 production in the serum of B6-DK mice are consistent with previous findings that TFH cells are preferentially expanded in B6-DK mice. Additional experiments, for example, with ICOS−/− mice, are needed to document the contribution of elevated ICOS+ CD4 cells to antitumor responses and delineate the potential link between ICOS+ CD4 cells, IL-21 production, and enhanced antitumor responses. Recent analysis of spatiotemporal dynamics of the tumor–immune interaction during tumor progression revealed that CXCL13 and IL-21 constitute the pivotal antitumor players, and increased numbers of TFH and B cells are strongly correlated with a positive prognosis (33). Promotion of effector T cells by IL-21 in a mouse melanoma model, and the efficacy of IL-21 treatment of patients with metastatic melanoma suggest a multifaceted role for IL-21, in addition to its potential contribution to antibody-mediated antitumor responses noted here (34, 35).
Here we find that an increase in the ratio of CD8 Teff to CD8 Treg (and no change in CD4 Treg) within tumors of B6-DK mice compared with those in B6-WT mice is associated with enhanced antitumor activity after tumor vaccination. Our observations are congruent with previous findings that increased ratios of CD8 T cells to FoxP3+CD4+ Treg are associated with favorable prognosis in the context of human and mouse cancers (36–38). Blockade of CD8 Treg-dependent inhibition may tip the balance within the tumor microenvironment in favor of the Teff compartment, resulting in enhanced antitumor immunity.
We also detected anti-MIF and anti–Ang-2 autoantibodies in some GVAX-treated tumor-bearing B6-DK mice, but not in B6-WT controls (Fig. 4C). These antibodies may exert antitumor effects through the inhibition of the Tie-2 pathway in macrophages and MMP-9 production (10). Interestingly, increases in both anti-MIF and anti–Ang-2 antibodies were also noted in response to treatment with GVAX and anti-CTLA-4 antibody therapy and correlated with increased tumor vasculopathy (10). Whether antibody activity present in IgG-enriched sera from B6-DK mice directly contributes to the antitumor response requires further experiments, including evaluation of the effects of absorption of purified IgG by tumor lysate. These considerations suggest that the disruption of normal immunoregulatory T cells may favor increased autoantibody responses that can target molecular elements that contribute to tumor growth and spread.
Here we identify a new immune phenotype of Qa-1–mutant mice. Genetic disruption of the inhibitory interaction between CD8 Treg and Qa-1+ TFH cells results in the induction of robust autoimmune antitumor responses driven by enhanced TFH cell helper activity and the upregulation of therapeutic antibody production by B cells. Our data indicate that the CD8+ Teff–Treg ratio in tumor lymphocyte infiltrates may represent a useful prognostic index for cancer development. These data also suggest that specific depletion of CD8 Treg represents a novel and potentially effective strategy for cancer treatment.
Disclosure of Potential Conflicts of Interest
No potential conflicts of interest were disclosed.
The Editor-in-Chief of Cancer Immunology Research is an author of this article. In keeping with the AACR's editorial policy, the paper was peer reviewed and an AACR journal editor not affiliated with Cancer Immunology Research rendered the decision concerning acceptability.
Conception and design: D.A. Alvarez Arias, H.-J. Kim, G. Dranoff, H. Cantor
Development of methodology: D.A. Alvarez Arias, H.-J. Kim, H. Cantor
Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): D.A. Alvarez Arias, H.-J. Kim, P. Zhou, T.A.W. Holderried, X. Wang, H. Cantor
Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): D.A. Alvarez Arias, H.-J. Kim, G. Dranoff, H. Cantor
Writing, review, and/or revision of the manuscript: D.A. Alvarez Arias, H.-J. Kim, G. Dranoff, H. Cantor
Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): D.A. Alvarez Arias, T.A.W. Holderried
Study supervision: H.-J. Kim, H. Cantor
This work was supported in part by NIH research grant AI 037562 and a gift from the LeRoy Schecter Research Foundation (to H. Cantor) and NRSA Fellowships (T32AI07386; to D. Alvarez Arias) and (T32CA070083; to H-J. Kim). T.A.W. Holderried is supported by the University of Düsseldorf (Düsseldorf, Germany).
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.
The authors thank Dr. Lisa Cameron (DFCI Microscopy Core) for help with image preparation and analysis, A. Angel for manuscript and figure preparation, and V. Garcia for technical assistance.
Note: Supplementary data for this article are available at Cancer Immunology Research Online (http://cancerimmunolres.aacrjournals.org/).
- Received August 13, 2013.
- Revision received November 12, 2013.
- Accepted December 9, 2013.
- ©2013 American Association for Cancer Research.