PD-L1 in tumor microenvironment mediates resistance to oncolytic immunotherapy

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Abstract

<p class="first" id="d10266594e260">Intralesional therapy with oncolytic viruses (OVs) leads to the activation of local and systemic immune pathways, which may present targets for further combinatorial therapies. Here, we used human tumor histocultures as well as syngeneic tumor models treated with Newcastle disease virus (NDV) to identify a range of immune targets upregulated with OV treatment. Despite tumor infiltration of effector T lymphocytes in response to NDV, there was ongoing inhibition through programmed death ligand 1 (PD-L1), acting as a mechanism of early and late adaptive immune resistance to the type I IFN response and T cell infiltration, respectively. Systemic therapeutic targeting of programmed cell death receptor 1 (PD-1) or PD-L1 in combination with intratumoral NDV resulted in the rejection of both treated and distant tumors. These findings have implications for the timing of PD-1/PD-L1 blockade in conjunction with OV therapy and highlight the importance of understanding the adaptive mechanisms of immune resistance to specific OVs for the rational design of combinatorial approaches using these agents. </p>

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The blockade of immune checkpoints in cancer immunotherapy.

Drew M Pardoll (2012)

Among the most promising approaches to activating therapeutic antitumour immunity is the blockade of immune checkpoints. Immune checkpoints refer to a plethora of inhibitory pathways hardwired into the immune system that are crucial for maintaining self-tolerance and modulating the duration and amplitude of physiological immune responses in peripheral tissues in order to minimize collateral tissue damage. It is now clear that tumours co-opt certain immune-checkpoint pathways as a major mechanism of immune resistance, particularly against T cells that are specific for tumour antigens. Because many of the immune checkpoints are initiated by ligand-receptor interactions, they can be readily blocked by antibodies or modulated by recombinant forms of ligands or receptors. Cytotoxic T-lymphocyte-associated antigen 4 (CTLA4) antibodies were the first of this class of immunotherapeutics to achieve US Food and Drug Administration (FDA) approval. Preliminary clinical findings with blockers of additional immune-checkpoint proteins, such as programmed cell death protein 1 (PD1), indicate broad and diverse opportunities to enhance antitumour immunity with the potential to produce durable clinical responses.

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Host type I IFN signals are required for antitumor CD8+ T cell responses through CD8α+ dendritic cells

Mercedes Fuertes, Aalok K. Kacha, Justin Kline … (2011)

Most tumors express antigens that can be recognized by T cells of the host immune system (Huang et al., 1994; Boon and Old, 1997). Despite the expression of antigens, tumors grow progressively and evade immunity. It has generally been assumed that immune evasion is a result of a failure to initiate an antitumor adaptive immune response. However, recent results have indicated that in many instances, spontaneous T cell responses against tumor antigens can be detected in both human cancer patients and in murine models, and that immune escape in those cases appears to occur through dominant inhibition by immunoregulatory pathways (Vesely et al., 2011). For example, high frequencies of CD8+ T cells specific for MelanA/MART-1, MAGE-10, and NY-Eso-1 have been detected in the blood of subsets of patients with metastatic melanoma (Pittet et al., 1999; Valmori et al., 2001; Mortarini et al., 2003; Peterson et al., 2003). Spontaneous antibody responses against a range of tumor-associated antigens have been previously described (Tan and Zhang, 2008). Antibody responses in early stage prostate cancer have been reported to be detected before PSA becomes detectably elevated (Wang et al., 2005). Moreover, we and others have shown that some human melanoma metastases contain activated CD8+ T cells, including tumor-reactive cells (Anichini et al., 1999; Harlin et al., 2009), suggesting that spontaneous immune responses can be generated all the way through to the step of effector cell migration into tumor sites. Expression of multiple immune evasion mechanisms likely blunts immune function at the effector phase and allows tumor outgrowth in those instances (Rabinovich et al., 2007). The observation that a T cell response can ever become spontaneously primed against a growing tumor mass raises the question of how this is possible given the tight regulation of innate immune signals that dictate whether a bridge to adaptive immunity can occur. Most malignancies (including melanoma) lack an obvious infectious etiology and therefore would not contain abundant external ligands for Toll-like receptors (TLRs). In this context, studies from several groups have revealed that dying cells can release endogenous adjuvants (Kono and Rock, 2008), providing activation signals for DCs and other APCs that lead to up-regulation of co-stimulatory molecules and consequently yield productive T cell activation and differentiation (Kono and Rock, 2008). Although these early results indicate that tumor cells can, under certain conditions, liberate products that can theoretically elicit innate immune signals, how these or other signals may lead to the spontaneous activation of a tumor-specific adaptive T cell response remains unclear. Type I IFNs have been studied extensively in the context of viral infections (Stetson and Medzhitov, 2006b). During various types of viral infection, type I IFNs induce the expression of an array of genes that act to prevent viral spread, thus creating an antiviral state (Stark et al., 1998). But type I IFNs also regulate antiviral immune effector responses and play an important role in promoting the cross-presentation of viral antigens to CD8+ T cells (Le Bon et al., 2003). Although a role for type I IFNs has been described for immunosurveillance against carcinogen-induced tumors and for rejection of transplanted tumors (Dunn et al., 2005, 2006), the source of type I IFNs and the mechanism of action of this cytokine during the priming phase of an antitumor immune response have not yet been elucidated. We have recently reported that gene expression profiling of human melanoma metastases revealed a subset of tumors that contained infiltrating CD8+ T cells (Harlin et al., 2009). Reasoning that interrogation of those gene array data might provide an indication regarding innate immune signals associated with the presence of a T cell response, we herein report a correlation between the presence of T cell–specific transcripts and a set of genes known to be induced by type I IFNs. Using a series of murine models, we show that shortly after tumor challenge in vivo, type I IFN production was detected by DCs in tumor-draining lymph nodes, and that host type I IFN signaling on CD8α+ DCs was required for spontaneous cross-priming of tumor antigen–specific CD8+ T cells. Our results suggest a model in which a growing tumor can elicit production of type I IFNs from the host, which leads to accumulation of CD8α+ DCs that in turn promote CD8+ T cell activation against tumor-derived antigens. RESULTS Tumors growing in immunocompetent hosts induce T cell priming and IFN-β production by CD11c+ cells in tumor-draining lymph nodes Gene expression profiling of human melanoma metastases along with confirmatory assays revealed that a subset of tumors showed evidence of spontaneous inflammation that included the presence of infiltrating CD8+ T cells (Harlin et al., 2009). Parallel studies have indicated that among tumor-associated CD8+ T cells there are cells that recognize human melanoma antigens as reflected by class I MHC/peptide tetramer staining (Speiser et al., 2002; Harlin et al., 2006). This evidence of spontaneous T cell priming against tumor antigens raised the question of what innate immune signals might mediate sterile immunity against tumors arising from self tissues. Reanalysis of the gene expression profiling data revealed that several transcripts indicative of type I IFN signaling, such as IRF1 and the IFN-induced protein of 30 kD were coexpressed in those tumors that contained T cell transcripts (Fig. 1). This correlation prompted mechanistic experiments to determine whether host type I IFN signals might be necessary for spontaneous priming of CD8+ T cells against tumor antigens when it does occur. Figure 1. Human melanoma metastases show a positive correlation between T cell markers and IFN-induced transcripts. Tumor samples were obtained by core biopsy or excisional biopsy or obtained from material resected from patients as part of routine clinical management. Total RNA was extracted from tumor samples (n = 52) and gene levels were analyzed by Affymetrix. Arbitrary expression units according to Affymetrix gene levels are shown. (IRF1, R2 = 0.648; p30, R2 = 0.658). We investigated whether spontaneous priming of antigen-specific CD8+ T cells could be detected in lymphoid organs after subcutaneous implantation of murine cell lines in immunocompetent mice. To provide a defined antigen, tumor cells were transduced to express the model antigenic peptide SIYRYYGL (SIY) that is presented by the class I molecule Kb. The SIY antigen is advantageous because of the set of tools we have assembled for monitoring details of the host immune response, including peptide/MHC tetramers, TCR transgenic T cells, and high-affinity TCR tetramers to monitor processed antigen on APCs (Kline et al., 2008; Zhang et al., 2008). B16 melanoma cells expressing the SIY antigen (B16.SIY) induced a significant frequency of peptide-specific IFN-γ–producing cells in the spleen 6 d after subcutaneous tumor implantation in the flank (Fig. 2 A). This was accompanied by an increase in the frequency of SIY-specific CD8+ T cells detected by SIY/Kb tetramer staining (Fig. 2 B). This T cell activation is completely dependent on cross-presentation by host DCs, as it was ablated in CD11c-DTR transgenic mice treated with diphtheria toxin (Fig. S1). Flow cytometry confirmed that the majority of CD8α+ and plasmacytoid DCs (pDCs), and a fraction of myeloid DCs (mDCs), were depleted with this approach (Fig. S1). It is interesting to note that these tumors grow progressively over time and are not ultimately rejected by the host, but this failure is not caused by an absence of early T cell priming. Rather, recent observations suggest that the immune response wanes over time because of dominant-negative regulatory mechanisms that eventually lead to tumor outgrowth (Kline et al., 2008). Regardless of those late events, early T cell priming can be used in this model as a readout for determining the host innate immune requirements for the initial recognition of tumor. Figure 2. Tumors induce CD8+ T cell priming, which is accompanied by IFN-β production by CD11c+ DCs in the tumor-draining lymph nodes. (A and B) C57BL/6 mice were inoculated or not with 5 × 106 B16.SIY tumor cells (s.c.), splenocytes were harvested 6 d later, and restimulated for 16 h in the presence or absence of soluble SIY peptide (A). The frequency of tumor-specific IFN-γ–producing cells was assessed by ELISPOT. ***, P < 0.0001 versus No tumor. (B) Cells were gated on CD8+CD4−B220−, and the frequency of SIY-specific CD8+ T cells was assessed by FACS using specific tetramers. **, P = 0.0063 versus No tumor. (C and D) C57BL/6 mice were inoculated (s.c.) or not with 5 × 106 B16.SIY tumor cells, and inguinal lymph nodes were recovered 4–6 d later. IFN-β mRNA expression was assessed by real-time RT-PCR analysis in total lymph nodes, and the results are expressed as 2−ΔCt using GAPDH as endogenous control. **, P = 0.0046 versus No tumor (C) or in CD11c+ and CD11c− cells sorted from lymph nodes. ***, P = 0.0008 versus CD11c− (D). (E) Wild-type C57BL/6 mice (expressing the congenic marker CD45.1+) were lethally irradiated and reconstituted with either wild-type (CD45.2+) or CD11c-DTR (CD45.2+) BM cells. Mice were allowed to reconstitute for 3 mo, and then were injected i.p. with diphtheria toxin (100 ng in 100 µl of DPBS) once a day for 8 d, starting 2 d before s.c challenge with 5 × 106 B16.SIY tumor cells in the left flank (n = 5). Inguinal lymph nodes were recovered 6 d later, and IFN-β expression in total inguinal lymph nodes was measured by real-time PCR. The results are expressed as 2−ΔCt using 18s as endogenous control. (F and G) C57BL/6 mice were inoculated s.c. with the indicated tumor cell lines (5 × 106; F), and splenocytes were harvested 6 d later and restimulated for 16 h in the presence or absence of soluble SIY peptide. Frequency of tumor-specific IFN-γ–producing cells was assessed by ELISPOT. (G) Real-time RT-PCR analysis of IFN-β mRNA expression in total inguinal lymph nodes. The results are expressed as 2−ΔCt using 18s as endogenous control. Data represent mean ± SEM (n = 5) and are representative of four independent experiments (A–C) or two independent experiments (D–G). To determine whether implanted tumors might also induce a type I IFN profile in mice, we inoculated B16 melanoma cells into recombination-activating gene 2-deficient (Rag2−/−) mice, to eliminate a contribution of the adaptive immune system. We compared gene expression of the tumors recovered from the mice to the B16 cells grown in vitro, reasoning that the differentially expressed genes would be those induced in the host by the presence of the tumor. Interestingly, this gene expression profiling confirmed the up-regulation of multiple transcripts reflective of innate immune activation, including a panel of IFN-inducible genes (Table S1). A repeat of this experiment in type I IFNR−/− mice confirmed that induction of these transcripts required type I IFN signaling on host cells (unpublished data). To assess directly whether type I IFNs are produced early in response to a growing tumor, we compared IFN-β mRNA levels by quantitative real-time PCR in lymph nodes from naive mice and from draining lymph nodes of C57BL/6 mice challenged with B16.SIY melanoma cells. We found that IFN-β was produced in tumor draining lymph nodes as early as 4 d after tumor challenge (Fig. 2 C). We next sorted cells from tumor draining lymph nodes on the basis of their expression of the DC marker CD11c and analyzed IFN-β mRNA levels, which revealed that IFN-β production after tumor challenge was confined to the CD11c+ DC subpopulation (Fig. 2 D). As an alternative approach to determining whether DCs were the dominant population producing IFN-β, we depleted DCs in vivo using CD11c-DTR transgenic mice treated with diphtheria toxin. After tumor implantation, these mice showed markedly reduced production of IFN-β in the tumor-draining lymph node compared with control mice (Fig. 2 E). Together, these results indicate that DCs are the major source of type I IFNs early after tumor challenge. To determine whether this ability to induce a rapid host immune response and type I IFN production was a general phenomenon, an additional panel of C57BL/6-derived tumor cells was transduced to express the SIY antigen. All the tumor lines tested (EL4, MC57, and C1498) similarly induced a spontaneous CD8+ T cell response as assessed by ELISPOT (Fig. 2 F) and peptide/MHC tetramer analysis (not depicted), and IFN-β production in the tumor-draining lymph node (Fig. 2 G). Together, these results indicated that it is not uncommon for implanted tumors to trigger type I IFN production and a rapid CD8+ T cell response against tumor-associated antigens in vivo. Host IFN-α/β signaling is critical for spontaneous tumor-specific T cell priming Having observed the early production of type I IFNs after tumor inoculation, we sought to determine whether host type I IFN signaling was necessary for spontaneous priming of CD8+ T cells to tumor antigens in vivo. We therefore examined the effect of B16.SIY challenge on the CD8+ T cell response to the SIY tumor antigen in mice deficient for the IFN-α/βR compared with WT mice. 17 d after tumor inoculation, splenocytes were assayed for the frequency of IFN-γ–producing cells in response to soluble SIY peptide or irradiated B16.SIY cells by ELISPOT. In comparison to the vigorous response observed in the wild-type group, splenocytes from IFN-α/βR−/− mice displayed a dramatically reduced frequency of IFN-γ–producing effector cells after tumor challenge (Fig. 3 A). To determine whether the effect of type I IFNs on T cell priming was at the level of expansion versus differentiation of CD8+ T cells, analysis using SIY/Kb tetramers was performed. Whereas an expanded population of tetramer-reactive CD8+ T cells was detected in the spleens of wild-type mice, IFN-α/βR−/−, and IFN-α/βR−/−/IFN-γR−/− mice failed to display an increased frequency of tumor reactive CD8+ T cells (Fig. 3 B). The combined elimination of IFN-α/βR and IFN-γR reduced T cell priming completely to background levels, suggesting partial compensation by IFN-γ in the absence of type I IFN signaling. However, mice singly deficient in the IFN-γR generated a normal expanded frequency of SIY-specific CD8+ T cells (Fig. 3 C), suggesting that type I IFNs are dominantly required and that IFN-γ is not necessary for this stage of an antitumor T cell response. Figure 3. IFN-α/β, and not IFN-γ signaling, is critical for spontaneous CD8+ T cell priming to tumor-associated antigens. (A and B) Wild-type, IFN-α/βR−/−, or IFN-α/βR−/−/IFN-γR−/− mice were inoculated s.c. with 106 B16.SIY tumor cells. Splenocytes were harvested 17 d later and restimulated for 16 h in the presence or absence of or soluble SIY peptide (A). The frequency of tumor-specific IFN-γ–producing cells was assessed by ELISPOT. ***, P < 0.0001 versus WT. (B) cells were gated on CD8+CD4−B220− and the frequency of SIY-specific CD8+ T cells was assessed by FACS using specific tetramers. ***, P < 0.0009; **, P < 0.0027 versus WT. (C) Wild-type and IFN-γR−/− mice were inoculated s.c. with 106 B16.SIY tumor cells, and splenocytes were harvested 17 d later and restimulated for 16 h in the presence or absence of soluble SIY peptide, and then the frequency of tumor-specific IFN-γ–producing cells was assessed by ELISPOT. P = 0.268 versus WT. (D and E) Wild-type and Stat1−/− mice were inoculated s.c. with 106 B16.SIY tumor cells, and splenocytes were harvested 17 d later and restimulated for 16 h in the presence or absence of soluble SIY peptide (D). The frequency of tumor-specific IFN-γ–producing cells was assessed by ELISPOT. ***, P < 0.0001 versus WT. (E) cells were gated on CD8+CD4−B220−, and the frequency of SIY-specific CD8+ T cells was assessed by FACS using specific tetramers. **, P = 0.0029 versus WT. Data represent mean ± SEM (n = 5), and are representative of three independent experiments. As type I IFNs mediate signaling through Stat1, we additionally analyzed spontaneous CD8+ T cell priming in Stat1−/− mice. Consistent with our previous results, Stat1−/− mice also showed a markedly decreased priming of tumor antigen–specific CD8+ T cells as assessed by ELISPOT and SIY/Kb tetramer analysis (Fig. 3, D and E). Together, these results indicate that host type I IFN signaling through Stat1 is required for optimal expansion of tumor antigen–specific CD8+ T cells after tumor challenge in vivo. IFN signaling is required in the hematopoietic compartment for spontaneous rejection of tumors in vivo To determine whether the defect in natural CD8+ T cell priming in the absence of host IFN signaling would be associated with failed tumor rejection, we used a model in which tumors are spontaneously rejected. At the same time, it was of interest to determine the cellular compartment in which IFN signaling must occur. Toward this end, we turned to the immunogenic variant of the P815 mastocytoma, P198, which initially grows but is spontaneously rejected in syngeneic DBA/2 mice (Fallarino et al., 1996). To determine if type I IFN signaling for tumor rejection was required in the hematopoietic or nonhematopoietic compartment, we generated radiation BM chimeras in which Stat1-sufficient versus Stat1−/− DBA/2 mice were irradiated and reconstituted with either wild-type or Stat1−/− DBA/2 BM cells. After a period of at least 3 mo, to allow for recovery of the immune system, mice received a tumor challenge with P198 cells on the flank and were monitored for tumor progression. As expected, P198 cells were rejected by wild-type mice that had been reconstituted with wild-type BM, and tumors grew progressively in Stat1−/− mice reconstituted with Stat1−/− BM (Fig. 4 A). In contrast, Stat1−/− mice reconstituted with wild-type BM cells rejected the tumor normally, indicating that the expression of Stat1 in hematopoietic cells is sufficient for tumor rejection and that Stat1 expression is not required in non-BM-derived cells for tumor immunity (Fig. 4 A). In addition, the failure of wild-type mice reconstituted with Stat1−/− BM to reject the P198 tumor demonstrates that expression of Stat1 in the hematopoietic compartment is necessary for spontaneous tumor rejection in vivo and that Stat1 in non-BM-derived cells is not sufficient for tumor rejection (Fig. 4 A). Figure 4. IFN signaling is required in non–T cell BMDCs for tumor-specific T cell priming and for spontaneous rejection of immunogenic tumors in vivo. (A) Wild-type and Stat1−/− DBA/2 mice were lethally irradiated and reconstituted with either wild-type or Stat1−/− DBA/2 BM cells, and 3 mo later they were challenged s.c. with 106 P198 cells in the left flank (n = 5). Tumor size was measured at different time points. Results are shown as mean tumor diameter ± SEM. Data are representative of two independent experiments. (B) T cells from wild-type and Stat1−/− (H-2b) mice were stimulated with T cell–depleted irradiated splenocytes from DBA/2 (H-2d) mice for 5 d in an allogeneic MLR. Cytotoxic activity was measured by standard 51Cr-release assay against P815 (H-2d, cognate targets) and EL4 cells (H-2b, control targets). (C and D) CD8+ T cells were purified from the spleens of 2C Tg/RAG2−/− (Stat1-sufficient) mice, CFSE labeled, and transferred by retroorbital injection to wild-type or Stat1−/− mice. The next day, those mice were inoculated s.c. with 106 B16.SIY cells in the flank, and 7 d later splenocytes were harvested and CFSE dilution of 2C CD8+ T cells was assessed by FACS. (C) Percent of cells with decreased CFSE intensity within the DAPI−CD8+1B2+ gate. Data show mean ± SEM of individual mice in each group. Data are representative of two independent experiments. (D) Representative dot plots of CFSE dilution. Numbers indicate percent of cells in the indicated gate. (E) 106 SIY-pulsed wild-type BMDCs were s.c. inoculated in the flank of wild-type or Stat1−/− mice, and 20 d later splenocytes were harvested and restimulated in vitro for 16 h in the presence of culture medium or soluble SIY peptide. The frequency of tumor-specific IFN-γ–producing cells was assessed by ELISPOT. Data are representative of two independent experiments. IFN signaling is required in non-T cell BM-derived cells for tumor antigen–specific T cell priming Among the cells of the hematopoietic compartment, IFN signaling could be playing a T cell–intrinsic role in the generation of an antitumor immune response and/or be important in non–T cells, perhaps at the level of APCs. To address whether T cell–intrinsic IFN signaling was required for the acquisition of an effector phenotype, we compared wild-type and Stat1−/− T cells primed to become effector CTL in a mixed lymphocyte culture in vitro. However, Stat1−/− and wild-type T cells were equally able to lyse allogeneic target cells (Fig. 4 B), suggesting that T cell–intrinsic Stat1 is not absolutely required for the development and execution of lytic effector function by cytotoxic T lymphocytes. IFN-γ production by Stat1−/− CTL was also preserved (unpublished data). To investigate more directly whether IFN signaling on non–T cells was required for activation of tumor antigen–specific CD8+ T cells, we CFSE labeled wild-type CD8+ T cells purified from 2C TCR Tg/Rag2−/− mice (Sha et al., 1988) specific for the SIY octameric peptide in the context of Kb (Udaka et al., 1996) and adoptively transferred them into wild-type or Stat1−/− syngeneic mice. We then challenged the mice with B16.SIY tumors. 7 d after tumor challenge with B16.SIY, we analyzed CFSE dilution of SIY-specific CD8+ T cells in the spleen. A large percentage of the 2C CD8+ T cells transferred into wild-type mice displayed a decreased intensity of CFSE fluorescence, consistent with antigen-specific T cell proliferation and successful cross-presentation of the SIY antigen. In contrast, CFSE dilution was markedly reduced upon transfer into Stat1−/− hosts (Fig. 4, C and D). Finally, to determine whether T cell priming could occur in the absence of T cell–intrinsic IFN signaling in vivo, we vaccinated Stat1−/− versus wild-type mice with wild-type BM-derived DCs (BMDCs) loaded with SIY peptide. When assayed 20 d later for T cell priming by IFN-γ ELISPOT, comparable induction of SIY-specific T cells was observed in both wild-type and Stat1−/− hosts (Fig. 4 E). This result demonstrates that T cell–intrinsic IFN signaling is not required for T cell priming in vivo, and suggests that the defect in T cell priming lies upstream, likely at the level of host APCs. Analysis of DC subsets from mice deficient in IFN signaling In a search for potential mechanisms responsible for the lack of tumor antigen–specific CD8+ T cell priming in the absence of host IFN signaling, we analyzed multiple phenotypic characteristics of DCs from wild-type versus Stat1−/− or IFN-α/βR−/− mice. We first compared frequencies and absolute numbers of the different DC subpopulations (mDCs, CD11C+B220−CD8α−CD11b+; CD8α+DCs, CD11C+B220−CD8α+CD11b−; and pDCs, CD11CintB220+PDCA+) in the spleen and lymph nodes of unmanipulated or tumor-bearing mice but found no difference between Stat1−/− or IFN-α/βR−/− and wild-type mice (unpublished data). In addition, the surface expression of CD80, CD86, CD40, and class I and class II MHCs in CD11c+ tumor-draining lymph node cells was comparable (Fig. 5 A and Table S2), and LPS-induced IL-12 production was still detected in Stat1−/− CD11c+ cells (not depicted). Moreover, adherent splenocytes from wild-type and Stat1−/− mice did not differ in their ability to stimulate 2C TCR Tg CD8+ T cells to produce IL-2 upon loading with the SIY peptide in vitro (Fig. 5 B). Figure 5. Analysis of DCs from WT, Stat1−/−, and IFN-α/βR−/− mice. (A) Wild-type and IFN-α/βR−/− mice were inoculated s.c. with 106 B16.SIY cells. 6 d later, surface expression of CD80, CD86, CD40, and class I and II MHC was assessed by FACS in tumor-draining lymph node cells gated on CD11c+ cells. Filled histograms correspond to an isotype control (IC), continuous line corresponds to wild-type, and dashed line corresponds to IFN-α/βR−/− mice. (B) Adherent splenocytes from wild-type and Stat1−/− mice were loaded with SIY peptide or left untreated and used to stimulate 2C CD8+ T cells. IL-2 production was assessed by ELISA. (C and D) Wild-type Rag1−/− mice (top) or Rag1−/−Stat1−/− mice (bottom) were inoculated with 106 MC57 or MC57.SIY tumor cells. 14 d later SIY/Kb expression was assessed by FACS using high-affinity 2C TCR tetramers gated on the tumor-infiltrating CD11c+ population (C) and CD11b+ population (D). Filled histograms correspond to staining with streptavidin-phycoerythrin alone. Data are representative of two independent experiments (n = 4). It was conceivable that host IFN signaling was generally important for the migration of DCs into the draining lymph node compartment in vivo. To address this possibility, the skin of wild-type and Stat1−/− mice was painted with FITC and the draining lymph nodes were analyzed for green fluorescence on CD11c+ cells. However, comparable appearance of FITC+ DCs was observed in both sets of mice (unpublished data). We also considered the possibility that type I IFNs would induce chemokines by the DCs that would aid in the recruitment of T cells to the lymph node for subsequent activation. Analysis of DCs stimulated with IFNα/β revealed substantial production of CXCL9 and CXCL10, which could potentially mediate recruitment of activated CD8+ T cells via CXCR3 (unpublished data). However, there was no defect in priming of SIY-specific CD8+ T cells in response to B16.SIY tumors in CXCR3−/− mice (unpublished data), arguing for a lack of importance of this pathway at the early stages of T cell priming in vivo. Finally, it was reasoned that type I IFN signaling might be necessary for DCs to be able to take up, process, and present tumor antigen in the context of class I MHC molecules. This possibility was addressed through the use of tetramers generated from a high-affinity variant of the 2C TCR, which can bind to SIY–Kb complexes with sufficient avidity to allow analysis by flow cytometry. Analysis of the APC compartments was performed in wild-type versus Stat1−/− mice crossed to a RAG2−/− background, to eliminate the possibility that host T cells might eliminate antigen-expressing APCs. In fact, when tumor-derived CD11c+ or CD11b+ cells were analyzed from wild-type versus Stat1−/− RAG2−/− mice bearing MC57.SIY tumors, comparable binding of the 2C TCR tetramer was observed (Fig. 5, C and D). Absence of binding to APCs derived from wild-type MC57 tumors confirmed specificity of tetramer binding only when the SIY antigen was present. These results suggest that the APC subsets that accumulate in the tumor site process and present the SIY tumor-derived antigen normally, even in the absence of host type I IFN signaling. In vivo, endogenous type I IFN signaling is required for intratumoral accumulation of the CD8α+ DC subset Given the comparable properties of DC subsets in the lymphoid organs of wild-type and Stat1−/− mice, and the intact expression of SIY–Kb peptide–MHC complexes on the total APC populations that are present within the tumor site, we next addressed whether there was a defect at the level of the specific DC subsets that accumulate within the tumor microenvironment. We therefore challenged wild-type and Stat1−/− mice with B16.SIY cells and evaluated the frequency and absolute number of the three major DC subpopulations accumulating in the tumor at day 16. We found that the mDC and pDC subpopulations accumulated in the tumors of both groups of mice. However, the CD8α+ DC population, which accounted for up to 20% of the DCs infiltrating the tumors in wild-type mice, was almost completely absent in the tumors grown in Stat1−/− mice (Fig. 6, A and B; and Fig. S2). A similar defect in CD8α+ DC accumulation was observed in IFN-α/βR−/− mice (Fig. 6 C). As an alternative quantitative approach to evaluate the presence of CD8α+ DCs, molecular markers were used. XCR1 is a chemokine receptor exclusively expressed by CD8α+ DCs (Dorner et al., 2009) and Batf 3, a transcription factor preferentially expressed by CD8α+ DCs and absolutely required for their development. Using quantitative RT-PCR to analyze transcript abundance in tumors analyzed ex vivo, we found that both transcripts were highly expressed in tumors grown in wild-type mice, yet severely reduced in tumors grown in IFN-α/βR−/− mice (Fig. 6, D and E). Therefore, host IFN signaling appeared to be required for intratumoral accumulation of CD8α+ DCs within the tumor microenvironment. Figure 6. Endogenous type I IFN signaling is required for intratumoral accumulation of CD8α+ DCs. (A and B) Wild-type and Stat1−/− mice were inoculated s.c. with 106 B16.SIY cells, and 15 d later tumors were harvested and frequency (A) and percentages (B) of CD8α+ DCs, mDCs, and pDCs infiltrating tumors were analyzed by FACS. GFP+DAPI+CD3+ cells were gated out, and the different DCs subpopulations were identified as follows: mDCs, CD11C+B220−CD8α−CD11b+; CD8α+DCs, CD11C+B220−CD8α+CD11b−; and pDCs, CD11CintB220+PDCA+. Results are shown as mean ± SEM of 3 independent experiments (n = 4). (C–E) Wild-type and IFN-α/βR−/− mice were inoculated s.c. with 106 B16.SIY cells, and 15 d later tumors were harvested and frequency of intratumoral CD8α+ DCs was assessed by FACS (C). (D and E) XCR1 mRNA expression (D) and Batf3 mRNA expression (E) were assessed by real-time RT-PCR analysis on tumor homogenates. The results are expressed as 2−ΔCt using 18s as endogenous control. Results are shown as mean ± SEM of 2 independent experiments (n = 5). CD8α+ DCs have been shown to be the most important DC population for cross-presentation of antigens to CD8+ T cells in the setting of viral infection (Belz et al., 2004). Recent work has indicated that mice lacking the transcriptional regulator, Batf3, have a specific deficiency in the development of the CD8α+ DC lineage (which includes a CD103+ DC subset; Hildner et al., 2008). These mice also show defective cross-priming of CD8+ T cells in response to viruses, and were defective in control of immunogenic tumors in vivo (Hildner et al., 2008). We therefore investigated whether these DC subpopulations were required at the level of priming of CD8+ T cells to tumor-derived antigens. Wild-type and Batf3−/− mice were challenged with B16.SIY melanoma cells, and splenocytes were assayed 6 d later for the frequency of IFN-γ–producing cells by IFN-γ ELISPOT and SIY/Kb tetramer staining. Indeed, Batf3−/− mice showed a dramatically reduced frequency of IFN-γ–producing effector cells as compared with wild-type mice (Fig. 7, A and B). The poor T cell priming was comparable to the level of deficiency observed in IFN-α/βR−/− mice (Fig. 3 A), as was the tumor growth rate, which was accelerated in Batf3−/− and IFN-α/βR−/− mice (Fig. S3, A and B). Moreover, parental B16 melanoma cells, without the SIY antigen, also grew faster in Batf3−/− mice compared with wild-type (Fig. S3 C), indicating that this is a critical pathway for cross-presentation of natural endogenous tumor antigens. To determine whether CD8α+ DCs were themselves required for type I IFN production in response to tumor implantation, induction of IFN-β mRNA was assessed in tumor-draining lymph nodes in wild-type versus Batf3−/− mice. However, IFN-β induction was comparable in both sets of mice (Fig. 7 C). Collectively, these findings demonstrate that the CD8α+ DC subpopulation is critical for the spontaneous priming of tumor antigen–specific CD8+ T cells in response to a growing tumor, downstream from host type IFN production. Figure 7. CD8α+ DCs are critical for antitumor CD8+ T cell priming. Wild-type and Batf3−/− mice were inoculated s.c. with 106 B16.SIY cells. 6 d later, splenocytes were harvested and restimulated for 16 h in the presence of culture medium or soluble SIY peptide (A). The frequency of tumor-specific IFN-γ–producing cells was assessed by ELISPOT. ***, P < 0.0001 versus WT. (B) the frequency of SIY-specific CD8+ T cells was assessed by FACS using specific anti–Kb-SIY tetramers, cells were gated on the CD8+CD4−B220− population. ***, P < 0.001 versus WT. (C) IFN-β mRNA expression was assessed by real-time RT-PCR analysis in total lymph nodes. The results are expressed as 2−ΔCt using 18s as endogenous control. Results are shown as mean ± SEM (n = 5) and are representative of at least two experiments. To determine whether the CD8α+ DC lineage itself must respond to type I IFNs, wild-type mice were irradiated and reconstituted with wild-type, Batf3−/− or IFN-α/βR−/− BM cells, or a mix of wild-type and IFN-α/βR−/− or Batf3−/− and IFN-α/βR−/− in a 50/50 ratio. 3 mo later, those chimeric mice were challenged with B16.SIY melanoma cells, and splenocytes were assayed 6 d later for T cell priming by SIY/Kb tetramer analysis and tumor size was measured. As expected, mice reconstituted with Batf3−/− or IFN-α/βR−/− BM cells showed a dramatically reduced frequency of SIY-specific CD8+ T cells as compared with mice reconstituted with wild-type BM cells (Fig. 8 A). Mice reconstituted with a mix of wild-type and IFN-α/βR−/− cells showed restored T cell priming to the level seen in wild-type mice. However, mice reconstituted with a mix of Batf3−/− and IFN-α/βR−/− BM cells continued to show reduced priming of SIY-specific T cells (Fig. 8 A). This reduced T cell priming was associated with poor tumor growth control (Fig. 8 B). These results demonstrate that type I IFNs must signal on the CD8α+ DC lineage for optimal priming of tumor antigen–specific CD8+ T cells after tumor challenge in vivo. Figure 8. Type I IFN signaling must occur on the CD8α+ DC lineage for antitumor CD8+ T cell priming to occur. (A) Wild-type mice were lethally irradiated and reconstituted with wild-type, Batf3−/−, or IFN-α/βR−/− BM cells, or a mix of wild-type and IFN-α/βR−/− or Batf3−/− and IFN-α/βR−/− in a 50/50 proportion. Mice were allowed to reconstitute for 3 mo, and were then inoculated s.c. with 106 B16.SIY cells. (A) splenocytes were harvested 6 d later, and the frequency of SIY-specific CD8+ T cells was assessed by FACS using specific tetramers. Cells were gated on CD8+CD4−B220−. (B) Tumor size was measured at the end of the experiment. *, P < 0.05 versus IFN-α/βR−/− + WT. Results are shown as mean ± SEM of 2 independent experiments (n = 4 each). DISCUSSION Our results identify type I IFNs as critical mediators in the spontaneous priming of an antitumor CD8+ T cell response. Our data show that IFN-β is produced shortly after tumor challenge and, through signaling at the level of CD8α+ DCs, promotes tumor antigen–specific T cell priming and tumor rejection. In vivo, endogenous type I IFNs induced the intratumoral accumulation of CD8α+ DCs, which were essential for spontaneous antitumor CD8+ T cell priming and which could explain the critical role of this cell lineage in spontaneous cross-priming of tumor antigen–specific CD8+ T cells. Given the lack of external TLR ligands for most malignancies, the molecular mechanism by which a tumor can provide the right environment to stimulate immune activation remains poorly understood. Emerging evidence indicates that dying cells can release endogenous adjuvants capable of activating APCs (Kono and Rock, 2008). Among these molecules are heat-shock proteins (Basu et al., 2000), uric acid (Shi et al., 2003), HMGB1 (high-mobility-group box 1; Scaffidi et al., 2002), ATP (Haag et al., 2007), and genomic double-stranded (ds) DNA (Ishii et al., 2001). It has been recently demonstrated that after radiotherapy or chemotherapy, dying tumor cells can release HMGB1 that binds to TLR4 (Apetoh et al., 2007) and ATP that activates the NALP3 inflammasome (Ghiringhelli et al., 2009). Presumably there is some degree of spontaneous tumor cell death either upon implantation of transplantable tumor cell lines or, as tumor growth exceeds the available blood supply during attempted neoangiogenesis. However, the massive tumor cell death occurring with chemotherapy or radiation is unlikely to be present in our system of spontaneous T cell priming, so other mechanisms could be operational. Recently, it has been shown that dsDNA can be released from necrotic cells, reach the cytoplasm of APCs, and be recognized by a TLR-independent pathway leading to IRF3 activation and to type I IFN production (Stetson and Medzhitov, 2006a). It is interesting to speculate that a TLR-independent cytosolic DNA recognition pathway might be involved in innate tumor recognition, IFN-β production, and spontaneous priming of tumor antigen–specific T cells. Our data have demonstrated that, shortly after tumor challenge, IFN-β is produced by CD11c+ DCs in tumor-draining lymph nodes. However, the identity of the specific subset of DCs producing type I IFNs in response to tumor growth remains unclear. The fact that IFN-β production is still observed in Batf3−/− mice strongly suggest that the CD8α+ DC subpopulation is not required for type I IFN production. Preliminary studies of depletion of pDCs using the anti-PDCA (Krug et al., 2004) mAb have revealed that T cell priming and IFN-β production appear to be intact (unpublished data). Thus, it may be that conventional mDCs are capable of this function. Nonetheless, our data are consistent with a model in which at least two different DC subpopulations collaborate for the induction of spontaneous antitumor T cell priming. In this model, one DC subpopulation (likely either mDCs or pDCs) would sense the presence of the tumor and produce type I IFNs, which through signaling on CD8α+ DCs would promote effective cross-priming of CD8+ T cells. Consistent with this model, it has been recently shown using quantitative proteomics that the CD8α+ DC subpopulation selectively lacks the receptors and signaling molecules (such as DAI [Takaoka et al., 2007] and Sting [Ishikawa et al., 2009]) required for the detection of nucleotides in the cytoplasm (Luber et al., 2010), so if this is indeed the pathway involved in type I IFN production to tumor, a non-CD8α+ DC subpopulation would need to be involved. Although several studies have suggested that CD8α+ DC distribution is restricted to lymphoid organs (Randolph et al., 2008) our results clearly indicate that CD8α+ DCs (defined as CD3−CD11c+B220−CD11b−CD8α+ cells) can infiltrate tumors. In agreement with our findings, it has been reported that CD8α+ DCs can infiltrate transplantable and spontaneous melanomas in B6 mice (Preynat-Seauve et al., 2006) and sarcomas in BALB/c mice treated with Flt3L and GM-CSF (Berhanu et al., 2006), and that such recruitment is associated with tumor rejection. Even though we found an augmented expression of XCR1 transcripts in tumors growing in wild-type hosts compared with type I IFN receptor–deficient mice, this difference could not be explained by a differential expression of its ligand, the chemokine XCL1, which was present in the tumor microenvironment in both hosts (unpublished data). The detailed mechanism by which type I IFNs induce the intratumoral accumulation of CD8α+ DCs will be a crucial area for future investigation. It is noteworthy that, under conditions in which spontaneous priming of antitumor CD8+ T cells was not occurring, we detected expression of processed SIY peptide–Kb complexes on the surface of several subsets of APCs. These results suggest that APC subtypes other than CD8α+ DCs are capable of processing antigen into the class I compartment. Similar results have been reported by others. Hans Schreiber’s laboratory has observed expression of processed tumor-derived antigen in tumor-infiltrating macrophages (Zhang et al., 2007). In addition, in the TRAMP model, tumor-infiltrating DCs have been suggested to express processed antigen and behave in a tolerogenic rather than an activating fashion (Anderson et al., 2007). Thus, although the CD8α+ DC subset is quantitatively superior at cross-presenting exogenous antigen into the class I compartment, it is likely that additional qualitative differences explain their ability to better initiate CD8+ T cell priming. Although it would have been ideal to characterize the DCs that had successfully processed antigen and then trafficked to the tumor-draining lymph node, we were unable to detect such cells with the TCR tetramer in the lymph node compartment, arguing that the presumably small number of cells is below the threshold of detection. It also should be pointed out that we studied DC subsets in the tumor microenvironment at relatively late time points, because in small tumors it simply wasn’t technically possible to detect them reliably. So we can only infer that a similar defect in CD8α+ DC accumulation is occurring at early times after tumor implantation. Nonetheless, our subsequent experiments solidified a requirement for CD8α+ DCs, and for type I IFN signaling on these cells, in order to attain spontaneous CD8+ T cell priming. It is currently unknown what dictates why tumors in some patients are capable of inducing spontaneous tumor-antigen specific T cell priming whereas others are not. Single nucleotide polymorphisms (SNPs) in different genes involved in the type I IFN pathway have been reported, including IFNAR (Muldoon et al., 2001) and Stat1 (Fortunato et al., 2008), that could affect levels of expression of the mature proteins, leading to variation in the response to type I IFNs. Alternatively, activation of distinct combinations of oncogenic pathways in individual tumors could lead to expression of distinct sets of genes that facilitate innate immune recognition in vivo. The involvement of type I IFNs in antitumor immune responses has been appreciated for a number of years. Although early clinical trials of systemic administration of type I IFNs showed encouraging results for the treatment of a broad range of tumors (Neidhart et al., 1984; Motzer et al., 2002), the mechanism by which exogenously administered type I IFNs induces antitumor activity has remained elusive. In addition, injection of mice with blocking antibody to IFN-α/β has been reported to enhance tumor growth, suggesting the importance of endogenous type I IFNs after tumor challenge and a role in inhibiting tumor growth in immunocompetent mice (Gresser et al., 1983). Our findings now describe a link between spontaneous IFN-β production and signaling on CD8α+ DC which is essential for tumor antigen–specific CD8+ T cell priming. In addition, preliminary data have revealed potent rejection of B16 melanoma when transduced to express IFN-β (unpublished data). Collectively, our results have implications for human cancer therapy, providing a strong rationale for the intratumoral administration of type I IFNs, which are already FDA approved for other indications, in order to promote improved activation of tumor antigen–specific CD8+ T cells using the tumor itself as a source of antigen. MATERIALS AND METHODS Human samples and gene array analysis. Biopsy processing and gene array analysis were described previously (Harlin et al., 2009). Data were interrogated for expression of IFN-regulated genes and referenced to TCR transcripts in individual tumors. Mice. C57BL/6 mice, 129 mice, and Stat1−/− mice were purchased from Taconic. For the indicated experiments, Stat1−/− mice were backcrossed for six generations onto DBA/2 mice (Jackson ImmunoResearch Laboratories). IFN-γR−/− mice were obtained from The Jackson Laboratory. IFN-αβR−/− mice and IFN-α/βR−/−IFN-γR−/− mice were purchased from B&K Universal. Batf3−/− mice (Hildner et al., 2008) were obtained from K. Murphy (Washington University School of Medicine, St. Louis, MO). Experiments in these strains were done either on the 129 or the C57BL/6 background (at least 5 generations) with similar results. 2C/RAG2−/− mice (Sha et al., 1988) were obtained from D. Loh (Washington University School of Medicine, St. Louis, MO). CD11c-DTR mice (Jung et al., 2002) were provided by D. Littman (New York University School of Medicine, New York, NY). All mice were used between 6 and 10 wk of age and were maintained in specific pathogen–free conditions in a barrier facility at the University of Chicago (Chicago, Illinois). All animal experiments were performed in accordance with protocol approved by the University of Chicago Institutional Animal Care and Use Committee. Tumor cell lines. The mutagenized DBA/2-derived mastocytoma cell line P198 and the C57BL/6 derived melanoma cell lines B16.F10 and B16.F10.SIY (henceforth referred to as B16.SIY), thymoma cell lines EL4 and EL4.SIY, fibrosarcoma cell lines MC-57 and MC-57.SIY, and the leukemia cell line C1498.SIY were used for experiments. All cells were maintained at 37°C with 7.5% CO2 in DME supplemented with 10% heat-inactivated FCS, MOPS, 2-mercaptoethanol, penicillin, streptomycin, l-arginine, l-glutamine, folic acid, and l-asparagine. In vivo tumor experiments. Cultured tumor cells were washed three times with Dulbecco’s PBS (DPBS), and 106 living cells were injected s.c. in 100 µl DPBS on the flank. For tumor growth experiments, the longest and shortest diameters were measured twice per week using calipers, and a mean and SD were calculated. For ELISPOT, tetramer staining, and CFSE dilution splenocytes were analyzed at the indicated time points after tumor challenge. For RT-PCR analysis of IFN-β, tumor-draining inguinal lymph nodes were collected and analyzed 4 to 6 d after tumor challenge. For analysis of tumor-infiltrating DC subpopulations, tumors were recovered 15 d after tumor challenge and were disrupted in complete DME medium containing 1 mg/ml collagenase IV (Sigma-Aldrich) for 30 min at 37°C. Data from groups of three to seven mice were analyzed. IFN-γ ELISPOT. The enzyme-linked Immunospot assay (ELISPOT) was conducted with the BD mouse IFN-γ kit according to the manufacturer’s protocol. Splenocytes were plated at 106 cells/well and stimulated overnight with irradiated (10,000 rad) B16.SIY cells (5 × 104/well), SIY peptide (80 nM), or PMA (50 ng/ml) and ionomycin (0.5 µM). In experiments analyzing priming by BMDCs, the restimulation was performed in FCS-free culture medium and plates were not blocked. IFN-γ spots were detected using biotinylated antibody and avidin-peroxidase and developed using AEC substrate (Sigma-Aldrich). Plates were read in an Immunospot Series 3 Analyzer and analyzed with ImmunoSpot software (Cellular Technology Ltd). Flow cytometry. Cells were incubated for 15 min at 4°C with anti-CD16 monoclonal antibody (2.4G2) to block potential nonspecific binding and with the fluorochrome-coupled antibodies against the following molecules for 30 min at 4°C: CD3ε, CD4, CD8, CD11b, CD11c, CD40, CD45R (B220), CD80, CD86, H-2Kb, and I-Ab/I-Eb (BD) and PDCA (Miltenyi Biotec). For tetramer staining, cells were labeled with PE-MHC class I tetramers (Beckman Coulter or Proimmune) consisting of murine H-2Kb complexed to either SIYRYYGL (SIY) peptide or SIINFEKL (OVA) peptide, anti–CD8-APC, anti–B220-PerCP-Cy5.5, and anti–CD4-PerCP-Cy5.5. For CFSE dilution analysis, splenocytes were labeled with DAPI, anti–CD8-APC, and 1B2-PE (anti–2C-TCR; Kranz et al., 1984). For TCR tetramer staining, cells were stained with anti–CD11c-APC, anti–CD11b-PerCp.Cy5.5, and biotin-2C-m67 TCR tetramer (Holler et al., 2003), followed by streptavidin-PE. FACS analysis was performed using FACSCanto or LSR II cytometers with FACSDiva software (BD). Data analysis was conducted with FlowJo software (Tree Star). For DC sorting experiments, pooled tumor-draining lymph node cells were stained with anti–CD11c-APC and sorted in a FACSAria cell sorter (BD). Quantitative real-time RT-PCR. Total RNA was purified using the RNeasy mini kit (QIAGEN) and analyzed in a 7300 Real Time PCR System (Applied Biosystems) using primer and probe sets from TaqMan Gene Expression Assays (Applied Biosystems) and TaqMan-based quantification. The results are expressed as 2−ΔCt using GAPDH or 18s as endogenous control. BM chimeras. Mice were maintained on trimethoprim and sulfamethoxazole for at least 3 d before the start of the experiment. Groups of 3–5 mice were lethally irradiated (900 rad) and maintained on antibiotics. The following day, pooled tibial and femoral BM cells from donor mice were ACK-lysed and 10 × 106 cells were injected retroorbitally into recipient mice. Mice were allowed to reconstitute for at least 8 wk before tumor challenge. Allogeneic MLR. MLR stimulation to generate effector cells for cytokine and CTL analysis was adopted from a previously published protocol (Gajewski et al., 1995). Total T cells were purified from spleens by negative selection with antibodies and magnetic beads from StemCell Technologies according to the manufacturer’s protocol. These responder cells were plated at 106/well containing stimulator cells consisting of allogeneic T cell–depleted irradiated (5,000 rad) splenocytes at 5 × 106/well. After 5 d, cells were used in chromium release assays and ELISA. Chromium release assay. Chromium release assays were performed as previously described (Kacha et al., 2000). Briefly, 51Cr-labeled targets (2 × 103) were plated with effectors cells at the indicated E/T ratios from 100:1 to 3.7:1. After 4 h of incubation at 37°C, 50 µl of supernatant was transferred to a LumaPlate-96 (PerkinElmer) and allowed to dry overnight. Plates were then counted using a TopCount-NXT plate reader (PerkinElmer). Percent specific lysis was calculated using standard methods. Cytokine ELISA. For cytokine analysis, tissue culture–treated 96-well flat bottom plates were coated with either DPBS alone, 2C11 (1 µg/ml; anti-CD3ε), or 2C11 and PV-1 (2 µg/ml; anti-CD28) in DPBS overnight at room temperature and washed with culture medium. Effector cells were incubated on the antibody-coated plates overnight, and supernatants were collected for measurement of IFN-γ concentration by ELISA. Mouse IL-2 and IFN-γ antibody sets were obtained from BD. Cytokine concentrations were determined with the Softmax PRO data analysis program (Molecular Devices). 2C CD8+ T cells purification, CFSE staining, and adoptive transfer. CD8+ T cells were purified from spleens of 2C/RAG2−/− mice by negative selection with antibodies and magnetic beads from StemCell Technologies according to the manufacturer’s protocol. T cells were stained with 2.5 mM CFSE at room temperature for 6 min and thoroughly washed with an excess volume of cold FCS. 107 CFSE-labeled T cells in 100 µl of DPBS were transferred by retroorbital injection into venous plexus of anesthetized mice. Mice were challenged with 106 B16.SIY cells, and 7 d later splenocytes from recipient mice were analyzed by flow cytometry. BMDC immunization. BMDCs were generated according to a modified version of a published protocol (Inaba et al., 1992). BM cells from the tibiae and femora were ACK-lysed and incubated for 10 d in complete DME medium with 20 ng/ml rmGM-CSF (R&D Systems) with the addition of 200 ng/ml LPS (Sigma-Aldrich) for the last 24 h. On day 10, cells were exposed to 10 µM SIY peptide for 1 h at 37°C and washed 3 times with DPBS. Mice were injected s.c. in the flank with 106 BMDCs in 100 µl DPBS. Adherent splenocyte stimulation of CD8+ T cells. Splenocytes were ACK-lysed, irradiated (5,000 rad), and plated at 106 cells/well. After 2 h of incubation, nonadherent cells were removed by 2 washes with DPBS. To stimulate CD8+ T cells, SIY peptide (10 µM) or culture medium was added to the adherent splenocytes, followed by the addition of 5 × 104 naive 2C/RAG2−/− CD8+ T cells/well. Supernatants were collected after 18 h and IL-2 production was determined by ELISA. Statistical methods. Differences between datasets were analyzed with the two-sided Student’s t test, and correlation was analyzed with Pearson test and Prism software (GraphPad). Online supplemental material. Fig. S1 shows that in CD11c-DTR transgenic mice treated with diphtheria toxin, the majority of CD8α+ DCs and pDCs, and a fraction of mDCs, were depleted and that these cell populations are critical for spontaneous CD8+ T cell priming to tumor-associated antigens. Fig. S2 shows that CD8α+ DCs fail to accumulate in the tumors grown in Stat1−/− mice. Fig. S3 shows that parental and SIY-expressing B16 melanomas grow faster in Batf3−/− and IFN-α/βR−/− mice compared with wild-type. Table S1 shows selected immune-related genes up-regulated with B16 tumors, including a panel of IFN-inducible genes. Table S2 shows that there is no difference in surface expression levels of CD40, CD80, and CD86 in CD11c+ tumor-draining lymph node cells from WT and IFN-α/βR−/− mice. Online supplemental material is available at http://www.jem.org/cgi/content/full/jem.20101159/DC1.

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Type I interferon is selectively required by dendritic cells for immune rejection of tumors

Mark Diamond, Michelle Kinder, Hirokazu Matsushita … (2011)

The ability of the immune system to function as an extrinsic tumor suppressor and effectively eliminate, control, and/or sculpt developing tumors forms the basis of the cancer immunoediting hypothesis (Shankaran et al., 2001; Dunn et al., 2002, 2004). There is strong experimental support for all three phases of cancer immunoediting, elimination, equilibrium, and escape, and many of the key cellular mediators and immune effector molecules involved in host protection from tumor development have been identified (Dunn et al., 2006; Smyth et al., 2006; Koebel et al., 2007; Schreiber et al., 2011; Vesely et al., 2011). The IFNs, both type I (IFN-α/β) and type II (IFN-γ), have emerged as critical components of the cancer immunoediting process, and work is ongoing to define their respective roles in promoting antitumor immune responses. Early studies supporting the existence of cancer immunoediting revealed an important function for IFN-γ in suppressing tumor development in models of both tumor transplantation and primary tumor induction (Dighe et al., 1994; Kaplan et al., 1998; Shankaran et al., 2001; Street et al., 2001, 2002). Specifically, IFN-γ was found to induce effects on both tumor cells (Dighe et al., 1994; Kaplan et al., 1998; Shankaran et al., 2001; Dunn et al., 2005) and host cells (Mumberg et al., 1999; Qin and Blankenstein, 2000; Qin et al., 2003). Subsequently, an essential function for endogenous type I IFN in cancer immunoediting was established (Dunn et al., 2005; Swann et al., 2007). Gene-targeted mice lacking the type I IFN receptor developed more carcinogen-induced primary tumors than WT control mice (Dunn et al., 2005; Swann et al., 2007), and antibody-mediated blockade of the IFN-α/β receptor in WT hosts abrogated rejection of immunogenic transplanted tumors (Dunn et al., 2005). The activity of endogenous type I IFN was mediated not by its direct effects on the tumor but by its actions on host cells, specifically on hematopoietic-derived host cells (Dunn et al., 2005). Collectively, these findings highlight a difference between the antitumor activities of the IFNs, wherein tumor cell responsiveness to IFN-γ but not IFN-α/β and host cell responsiveness to both IFN-γ and IFN-α/β are crucial for tumor rejection. However, the relevant host cell targets and antitumor functions of IFN-α/β and IFN-γ remain undefined because of the nearly ubiquitous expression of IFN-α/β and IFN-γ receptors and the pleiotropic effects they induce. Although initially defined by their antiviral activity, the type I IFNs are potent immunomodulators that shape host immunity through direct actions on innate and adaptive lymphocytes. The enhancement of NK cell cytotoxicity by IFN-α/β in the setting of viral infection was one of the earliest such effects to be recognized (Biron et al., 1999). Type I IFN directly augments NK cell–mediated killing of virally infected or transformed cells and indirectly promotes the expansion and survival of NK cells through IL-15 induction (Nguyen et al., 2002). Furthermore, in models of NK cell–dependent tumor rejection, host cell responsiveness to IFN-α/β was shown to be important for control of tumor growth and metastasis (Swann et al., 2007). Type I IFN can also act directly on T and B lymphocytes to modulate their activity and/or survival. Treatment with IFN-α/β in vitro prolonged the survival of activated T cells (Marrack et al., 1999) and augmented clonal expansion and effector differentiation of CD8+ T cells (Curtsinger et al., 2005) through cell-intrinsic IFN-α/β receptor signaling. Similarly, type I IFN responsiveness in T cells was required in vivo for optimal clonal expansion of antigen-specific CD8+ and CD4+ T cells during viral infection (Kolumam et al., 2005; Havenar-Daughton et al., 2006; Thompson et al., 2006) as well as for CD8+ T cell priming after immunization with antigen and IFN-α (Le Bon et al., 2006a). B cell differentiation, antibody production, and isotype class switching were also enhanced by type I IFN’s effects either directly on B cells or indirectly via effects on T cells (Coro et al., 2006; Le Bon et al., 2006b) and DCs (Le Bon et al., 2001). Type I IFN also directly enhances the function of DCs, which are central to the initiation of adaptive immune responses (Steinman and Banchereau, 2007). IFN-α/β induces DC maturation, up-regulates their co-stimulatory activity and enhances their capacity to present or cross-present antigen (Luft et al., 1998; Gallucci et al., 1999; Montoya et al., 2002). For example, coinjection of IFN-α/β plus antigen (Gallucci et al., 1999; Le Bon et al., 2001, 2003) or injection of DC-targeted antigen in combination with the IFN-α/β inducer polyinosinic:polycytidylic acid (polyI:C; Longhi et al., 2009) stimulated CD8+ T cell priming, humoral responses, and development of CD4+ Th1 responses in vivo. Recently, a subpopulation of DCs whose development depends on expression of the BATF3 transcription factor (CD8α+ DCs and CD103+ DCs, hereafter referred to as CD8α+ lineage DCs) was shown to play an important role in cross-presenting viral and tumor antigens, and mice lacking these cells fail to reject highly immunogenic unedited sarcomas (Hildner et al., 2008; Edelson et al., 2010). However, it remains unknown whether the cross-presenting activity of these cells requires type I IFN to induce tumor immunity. In the current study, we have investigated the host cell targets of endogenous type I IFN during the rejection of highly immunogenic, unedited tumors. We demonstrate that IFN-α/β acts early during the initiation of the immune response and that innate immune cells represent the essential responsive cells for the generation of protective antitumor immunity. Whereas type I IFN–unresponsive mice showed a defect in the priming of tumor-specific CTLs, reconstitution of IFN-α/β sensitivity in innate immune cells was sufficient to restore this deficit and resulted in tumor rejection. Within the innate immune compartment, we find no evidence of an essential role for NK cells or for type I IFN sensitivity in granulocytes or macrophages, but rather find that the actions of IFN-α/β on DCs are required for development of tumor immunity in vivo and play an important role in promoting the capacity of CD8α+ lineage DCs to cross-present antigen to CD8+ T cells. These results thus identify DCs and specifically CD8α+ lineage DCs as key cellular targets of type I IFN in the development of protective adaptive immune responses to immunogenic tumors. RESULTS Early requirement for type I IFN during the antitumor response We previously showed that blockade of type I IFN signaling by pretreatment of mice with the IFNAR1 (IFN-α/β receptor 1)-specific MAR1-5A3 mAb (Sheehan et al., 2006) abrogated rejection of highly immunogenic sarcomas derived from 3′-methylcholanthrene (MCA)–treated Rag2−/− mice (termed unedited tumors; Dunn et al., 2005). To dissect the temporal requirements for IFN-α/β’s actions during the antitumor immune response, we treated WT mice with either MAR1-5A3 or isotype control GIR-208 mAb at different times after injection of unedited H31m1 MCA sarcoma cells. Whereas H31m1 cells were rejected when transplanted into naive syngeneic WT mice either left untreated or pretreated with GIR-208, the tumors grew progressively in WT mice pretreated with MAR1-5A3 (Fig. 1 A). Similarly, MAR1-5A3 treatment on day 4 or 6 (relative to tumor injection at day 0) blocked rejection in >50% of injected mice. In contrast, IFN-α/β receptor blockade at later time points did not inhibit rejection (Fig. 1 B and Fig. S1). In parallel experiments, blockade of IFN-γ via treatment with neutralizing IFN-γ–specific H22 mAb (Schreiber et al., 1985) revealed a more prolonged requirement for the actions of IFN-γ during H31m1 rejection (Fig. 1 C). Cohorts of mice were also treated with a mixture of mAbs that deplete CD4+ and CD8+ cells and neutralize IFN-γ (GK1.5 [Dialynas et al., 1983], YTS169.4 [Cobbold et al., 1984], and H22, respectively) to broadly disrupt host immunity. In this group, progressively growing tumors were observed in a substantial proportion of mice treated as late as day 14 with the anti-CD4/CD8/IFN-γ mAb cocktail (Fig. 1 D). Collectively, these data demonstrate that the obligate functions of type I IFN are required only for initiating the immune response to tumors. Figure 1. Early requirement for IFN-α/β during rejection of highly immunogenic tumor cells. (A) Untreated WT and Rag2−/− mice or WT mice injected i.p. with either IFNAR1-specific MAR1-5A3 mAb or isotype control GIR-208 mAb 1 d prior were s.c. injected with 106 H31m1 tumor cells, and tumor size was measured over time. Data represent mean tumor diameter ± SEM of 12–16 mice per group from at least three independent experiments. (B–D) WT mice were injected with 106 H31m1 cells (at day 0) and treated beginning on the indicated day with MAR1-5A3 (B), IFN-γ–specific H22 mAb (C), or a mixture of anti-CD4/anti-CD8/anti–IFN-γ mAbs GK1.5/YTS-169.4/H22 (D), and tumor growth was monitored. For each time point, groups of mice were treated in parallel with the respective isotype-matched control mAb, and the data are presented as percent tumor growth over the control group. Results are from two to four experiments with 14–20 (ctrl/MAR1-5A3), 10–20 (ctrl/H22), or 6–11 (ctrl/cocktail) WT mice per group. The kinetics of tumor growth in individual mice is shown in Fig. S1. A tissue-restricted role for type I IFN during tumor rejection To characterize the critical host cells responding to type I IFN during initiation of the antitumor response, we transplanted H31m1 tumor cells and cells from a second unedited MCA sarcoma, d38m2, into bone marrow chimeras with selective IFN-α/β sensitivity. These tumor cell lines were selected because we previously showed that their rejection required type I IFN responsiveness at the level of the host (Dunn et al., 2005). As reported previously, both cell lines were rejected when transplanted into immunocompetent WT mice but formed progressively growing tumors in either Rag2−/− or Ifnar1−/− mice (Fig. 2, A and B). We now show that both lines grew progressively in Ifnar1−/− → Ifnar1−/− bone marrow chimeras and Ifnar1−/− → Rag2−/− chimeras (IFN-α/β sensitivity only in nonhematopoietic cells) but were rejected in WT → WT chimeras and WT → Ifnar1−/− chimeras (IFN-α/β sensitivity only in hematopoietic cells). These results thus extend, to two additional tumors, our prior finding that type I IFN sensitivity within the hematopoietic compartment is both necessary and sufficient for tumor rejection (Dunn et al., 2005). Figure 2. Nonoverlapping host cell targets for IFN-α/β and IFN-γ during tumor rejection. (A–C) Control mice and the indicated bone marrow chimeras with selective IFN-α/β sensitivity (A and B) or IFN-γ sensitivity (C) in hematopoietic versus nonhematopoietic cells were injected s.c. with 106 H31m1 (A) or d38m2 (B and C) unedited MCA sarcoma cells, and growth was monitored. Data are presented as mean tumor diameter ± SEM over time or the percentage of tumor-positive mice per group from two to three (A and B) or five (C) independent experiments with group sizes as indicated. Hematopoietic reconstitution of all Ifnar1−/− and Ifngr1−/− bone marrow chimeras was confirmed by flow cytometry at the conclusion of each experiment. Because the rejection of immunogenic sarcomas also requires IFN-γ sensitivity within the host (Fig. S2), we wanted to determine whether IFN-α/β and IFN-γ were mediating their effects by acting on the same host cell compartment. We thus performed a similar set of experiments using chimeras with selective host cell IFN-γ responsiveness. As expected, d38m2 tumor cells grew progressively in Rag2−/− , Ifngr1−/− , and Ifngr1−/− → Ifngr1−/− mice but were rejected in WT and WT → WT hosts (Fig. 2 C). Tumor growth was also observed in a significant fraction of Ifngr1−/− → Rag2−/− and WT → Ifngr1−/− chimeras, though the defect in these mice (which selectively express the IFN-γ receptor in either nonhematopoietic or hematopoietic cells, respectively) appeared less severe than that in globally insensitive Ifngr1−/− → Ifngr1−/− chimeras. To ensure that tumor growth in the chimeric mice was not caused by incomplete hematopoietic reconstitution, we confirmed normal cellularity and immune cell percentages in the spleen, demonstrated normal functional immune reconstitution, and ruled out the presence of radio-resistant tissue-resident leukocytes within the tumor environment (Figs. S3–S5). These data not only establish an important role for IFN-γ sensitivity in both hematopoietic and nonhematopoietic cells during tumor rejection but also reveal a difference between the broad cellular requirements for IFN-γ as opposed to the tissue-restricted requirement for IFN-α/β during elimination of the same tumor. Innate immune cells are the critical targets of type I IFN To examine the role of type I IFN’s actions on innate versus adaptive immune cells, we generated mixed bone marrow chimeras with selective type I IFN sensitivity within the hematopoietic compartment. Reconstitution of lethally irradiated Ifnar1−/− mice with a 4:1 mixture of Rag2−/− and Ifnar1−/− hematopoietic stem cells (HSCs) yielded mice with IFN-α/β responsiveness solely in innate immune cells (Rag2−/− + Ifnar1−/− → Ifnar1−/− chimeras, hereafter referred to as innate chimeras). Conversely, reconstitution of Ifnar1−/− mice with a 4:1 mixture of Rag2−/− × Ifnar1−/− double KO mice (Rag2−/−Ifnar1−/− ) and WT HSCs produced chimeras with IFN-α/β–sensitive T and B lymphocytes but a predominantly IFN-α/β–insensitive innate immune compartment (Rag2−/−Ifnar1−/− + WT → Ifnar1−/− ; adaptive chimeras). Control chimeras with responsiveness in both innate and adaptive compartments (Rag2−/− + WT → Ifnar1−/− ; innate + adaptive) or neither compartment (Ifnar1−/− → Ifnar1−/− ; “neither”) were also generated. The phenotypes of mixed chimeras generated using this approach were confirmed by IFNAR1 staining of splenocyte subsets (Fig. 3 A and Fig. S6). Figure 3. IFN-α/β sensitivity within the innate immune compartment is necessary and sufficient for tumor rejection. Mixed bone marrow chimeras with selective IFNAR1 expression in innate or adaptive immune cells were generated by reconstitution of irradiated Ifnar1−/− mice with mixtures of HSCs as described in Results. (A) Splenocytes were isolated from representative cohorts of control and mixed chimeric mice at least 12 wk after reconstitution, and IFNAR1 staining was analyzed by flow cytometry. Shown are the percentages of IFNAR1+ cells within the indicated immune cell subsets for 8–14 mice of each type. Horizontal bars represent the mean. (B–D) Control WT, Rag2−/− , and Ifnar1−/− mice and Ifnar1−/− mixed chimeric mice were injected with 106 H31m1 (B), d38m2 (C), or F515 (D) tumor cells, and growth was monitored over time. Data are presented as mean tumor diameter ± SEM or the percentage or tumor-positive mice per group from two to three independent experiments with group sizes as indicated. WT mice treated with control or IFN-γ–specific mAb were challenged with 106 F515 tumor cells, and growth was monitored (D, bottom). Mean tumor diameter ± SEM for 7–10 mice/group from two experiments is shown. When challenged with H31m1 or d38m2 cells, Rag2−/− and Ifnar1−/− control mice and globally unresponsive “neither” chimeras developed progressively growing tumors. In contrast, WT controls and pan-hematopoietic responsive innate + adaptive or WT → WT chimeras rejected the tumor challenge (Fig. 3, B and C), consistent with our previous results (Fig. 2). Importantly, H31m1 and d38m2 cells were rejected in mixed chimeras with IFN-α/β sensitivity only in innate immune cells (i.e., innate chimeras) but grew progressively in chimeras with IFN-α/β sensitivity largely restricted to the adaptive immune compartment (i.e., adaptive chimeras). These findings demonstrate that the essential antitumor functions of type I IFN on host cells during tumor rejection are selectively directed toward cells of the innate immune compartment. To confirm the functional hematopoietic reconstitution of Ifnar1−/− mixed chimeras, we performed three experiments. First, we confirmed the normal representation of various immune cell subsets within the spleens of mixed chimeric mice (Fig. 4, A and B). Second, we assessed the in vivo growth behavior of unedited MCA sarcoma cells (F515) that require lymphocytes and IFN-γ but not host IFN-α/β responsiveness for their rejection. F515 tumor cells grew progressively when injected into Rag2−/− mice and WT mice treated with IFN-γ–specific H22 mAb but were rejected in WT mice, WT mice treated with isotype control PIP mAb, and Ifnar1−/− hosts (Fig. 3 D). Similar to Ifnar1−/− mice, F515 cells were also rejected in Ifnar1−/− mixed chimeras of each type, verifying functional reconstitution of the immune compartment. Third, to rule out a potential hyperactive immunological state in these reconstituted mice, we challenged Ifnar1−/− mixed chimeras and control mice with MCA sarcoma cells derived from WT mice (1877). We have previously established that this tumor grows progressively when transplanted into naive WT mice (unpublished data). Similarly, these tumor cells grew progressively in Ifnar1−/− mixed chimeras of each type (Fig. 4 C). Figure 4. Normal hematopoietic reconstitution in Ifnar1−/− mixed bone marrow chimeras. (A) Spleens were harvested from WT, Ifnar1−/− , or Ifnar1−/− mixed chimeras of each type (12 wk after reconstitution), and cell density was determined. Horizontal bars represent the mean. (B) Percentages of the indicated immune cell subsets were measured by flow cytometry for WT, Ifnar1−/− , and Ifnar1−/− mixed chimeras. Mean values (as a percentage of total splenocytes) ± SEM for four to five mice/group are shown. Cell populations were defined as follows: CD4+ T cells (CD3+CD4+), CD8+ T cells (CD3+CD8+), B cells (B220+), NK cells (DX5+CD3−), DCs (CD11chi), and myeloid cells (CD11b+). (C) WT-derived 1877 tumor cells were injected at a dose of 106 cells/mouse into WT, Ifnar1−/− , Rag2−/− , and Ifnar1−/− mixed chimeras, and tumor growth was monitored over time. Data represent the mean tumor diameter ± SEM for three to eight mice/group. (A–C) Data are representative of two independent experiments. Sensitivity to type I IFN in innate immune cells is required for the generation of tumor-specific CTL To investigate the mechanism by which endogenous type I IFN promoted host antitumor responses, we looked specifically at the priming of tumor-specific T cells in WT and Ifnar1−/− mice after tumor challenge. Splenocytes from WT hosts isolated 20 d after inoculation of H31m1 tumor cells showed robust cytolytic activity against H31m1 targets after in vitro restimulation (Fig. 5 A). In contrast, tumor-specific killing was largely absent from splenocytes derived from Ifnar1−/− mice challenged with tumor cells. Similar results were observed using another highly immunogenic unedited MCA sarcoma (GAR4.GR1) or using IFN-γ production as a readout (unpublished data). To ask whether type I IFN sensitivity in innate immune cells was sufficient to generate tumor-specific immune responses, we used the mixed bone marrow chimeras described previously (Fig. 3). These experiments showed that IFN-α/β’s actions on the innate immune compartment were indeed both necessary and sufficient for development of tumor-specific cytotoxicity (Fig. 5 B). In addition, treatment of splenocytes from innate chimeras with blocking CD4- or CD8-specific antibodies confirmed the importance of CD8+ cells for in vitro cytotoxicity (Fig. 5 C). These results demonstrate the selective importance of type I IFN on innate immune cells to induce tumor-specific CTL priming. Figure 5. Impaired tumor-specific CTL priming in Ifnar1−/− mice is restored by IFN-α/β–responsive innate immune cells. (A) Splenocytes from WT and Ifnar1−/− mice were isolated 20 d after H31m1 tumor challenge (106 cells/mouse), co-cultured with IFN-γ–treated, irradiated H31m1 cells, and 5 d later used as effectors in a cytotoxicity assay with 51Cr-labeled H31m1 targets. Specific killing activity (in percentage ± SEM) at the indicated effector/target (E:T) ratios is shown for four to five mice per group assayed in duplicate from three independent experiments. (B) Splenocytes were harvested from the indicated chimeric mice 20 d after injection of 106 H31m1 tumor cells and were treated as in A. Data include representative results from three mice per group assayed in duplicate from two independent experiments. Splenocytes harvested from a naive mouse and treated similarly served as a negative control. (C) Effector cells from H31m1-challenged innate chimeras were co-cultured at the indicated effector/target ratios with 51Cr-labeled H31m1 targets in the presence of 10 µg/ml control (PIP), anti-CD4 (GK1.5), or anti-CD8 (YTS-169.4) mAbs. Data show representative results from three mice per group assayed in duplicate from three independent experiments. Similar results were obtained when effector cells from H31m1-injected WT mice were used (not depicted). (B and C) Error bars represent SEM. NK cells are not required for type I IFN–dependent tumor rejection Because NK cells have a host-protective function in some tumor models and display enhanced cytotoxic activity in response to type I IFN, we investigated the role of NK cells in the rejection of highly immunogenic sarcomas. We used comparable unedited MCA sarcoma cells generated from genetically pure C57BL/6 Rag2−/− mice and naive WT C57BL/6 mice as recipients because we could deplete NK cells in C57BL/6 mice with the NK1.1-specific PK136 mAb (Koo and Peppard, 1984). Similar to results with unedited MCA sarcomas from 129/Sv mice, immune-mediated rejection of two representative C57BL/6 strain unedited sarcomas (1969 and 7835) required IFN-α/β sensitivity at the level of the host (Fig. 6, A and B). When PK136-treated WT mice were injected with unedited C57BL/6 tumor cells, they rejected these tumors with kinetics identical to control mice. We confirmed NK cell depletion by (a) flow cytometry, (b) the absence of ex vivo killing of YAC-1 targets by splenocytes from mAb-treated mice, and (c) the lack of in vivo control of RMA-S tumor cell growth (Fig. 6, C–E). These data therefore indicate that NK1.1+ NK cells are not required for IFN-α/β–dependent rejection of unedited MCA sarcomas. Figure 6. NK cell depletion does not abrogate IFN-α/β–dependent rejection of immunogenic sarcomas. (A and B) C57BL/6 WT, Rag2−/− , and Ifnar1−/− mice and WT mice treated with either PBS or anti-NK1.1 PK136 mAb were injected s.c. (106 cells/mouse) with 1969 (A) or 7835 (B) unedited MCA sarcoma cells, and growth was monitored over time. Data are presented as mean tumor diameter ± SEM of 4–13 (untreated) or 8 (treated) mice per group from at least two independent experiments. Error bars for Ifnar1−/− mice reflect progressive growth of 1969 and 7835 tumors in 6/9 mice. (C) WT C57BL/6 mice were treated with either PBS or PK136 mAb, and splenocytes were harvested 2 d later and analyzed by flow cytometry using the NK cell markers DX5 and NKp46. Splenocytes were gated on CD3− cells, and the percentages of DX5+NKp46+ cells are indicated. Similar results were found when harvested at day 6 (not depicted). (D) WT C57BL/6 mice were treated with PBS or PK136 followed by i.p. injection of 300 µg polyI:C 4 d later. After 24 h, splenocytes were harvested and used as effectors in a standard 4-h cytotoxicity assay with NK-sensitive YAC-1 targets. Specific lysis (in percentage ± SEM) at the indicated effector/target (E:T) ratios is shown for four mice/group assayed in duplicate from two independent experiments. (E) WT C57BL/6 mice were treated with PBS, PK136, or a mixture of anti-CD4 (GK1.5) and anti-CD8 (YTS-169.4) mAbs and injected s.c. with 105 RMA-S cells, and tumor growth was monitored over time. Mean tumor diameter ± SEM for three mice/group is shown, and data are representative of two independent experiments. Granulocytes and macrophages do not require type I IFN sensitivity for tumor rejection To test whether type I IFN sensitivity is required by granulocytes and macrophages for tumor rejection, we crossed C57BL/6 strain LysM-Cre+ mice (Clausen et al., 1999) to C57BL/6 Ifnar1f/f mice (Prinz et al., 2008; prepared by backcrossing 129 strain Ifnar1f/f mice >99% onto a C57BL/6 background using a speed congenic approach). The resulting LysM-Cre+Ifnar1f/f mice displayed complete IFNAR1 deletion in peritoneal macrophages and PMNs and substantial deletion of IFNAR1 in monocytes (66%) and splenic macrophages (35%) but maintained undiminished IFNAR1 expression in DCs, NK cells, T cells, and B cells (Fig. 7, A and B). Peritoneal macrophages from these mice were unresponsive to type I IFN and failed to phosphorylate STAT1 after IFN-α stimulation (Fig. 7 C). However, LysM-Cre+Ifnar1f/f mice still rejected highly immunogenic unedited B6 strain 1969 sarcoma cells similar to IFN-α/β–responsive Ifnar1f/f mice (Fig. 7 D). In contrast, these tumor cells formed progressively growing tumors in B6 strain Ifnar1−/− control mice. Thus, protective tumor immunity does not require type I IFN sensitivity in granulocytes and at least some macrophage compartments. Figure 7. Granulocytes and macrophages do not require type I IFN sensitivity for tumor rejection. (A) IFNAR1 expression on peritoneal macrophages, blood monocytes, PMNs, and B cells was measured using flow cytometry in Ifnar1f/f , LysM-Cre+Ifnar1f/f , and Ifnar1−/− mice. (B) Summary of IFNAR1 levels in the indicated cellular subsets in LysM-Cre+Ifnar1f/f mice compared with Ifnar1f/f mice (expressed as a percentage of the mean fluorescence intensity [MFI]). Cells were gated using the following markers: macrophages (CD11b+F4/80+), PMNs (CD11b+Gr1+), monocytes (CD115+CD11b+), B cells (B220+), CD8α+ DCs (CD8α+Dec205+CD11chi), CD4+ DCs (CD8α−Dec205−CD11chiCD4+), pDCs (B220+PDCA+CD11cint), T cells (CD3+), and NK cells (NK1.1+). IFNAR1 expression was measured using MAR1-5A3 mAb. Data represent at least three mice from three independent experiments (**, P 99.7% purity). 129/Sv Rag2−/−Ifnar1−/− mice were generated by intercrossing Rag2−/− and Ifnar1−/− mice. OT-I transgenic mice on a Rag1−/− background were obtained through the National Institute of Allergy and Infectious Diseases Exchange Program, National Institutes of Health (C57BL6-Tg(OT-I)-RAG1tm1Mom 004175; Mombaerts et al., 1992; Hogquist et al., 1994). C57BL/6 MHC class I–deficient Kb−/−Db−/−β2m−/− mice (Lybarger et al., 2003) were a gift from H. Virgin and T. Hansen (Washington University in St. Louis, St. Louis, MO). 129/SvEv background Batf3−/− mice have been described previously (Hildner et al., 2008). Mice were maintained in a specific pathogen-free facility in accordance with American Association for Laboratory Animal Science guidelines, and all protocols involving laboratory animals were approved by the Washington University Animal Studies Committee (School of Medicine, Washington University in St. Louis). Tumor transplantation. MCA-induced fibrosarcomas were derived from 129/Sv strain Rag2−/− or WT mice and C57BL/6 strain Rag2−/− mice as described previously (Shankaran et al., 2001; Dunn et al., 2005; Koebel et al., 2007). The GAR4 tumor, derived from an MCA-treated 129/Sv Ifngr1−/−Ifnar1−/− mouse, as well as IFNGR1-resconstituted GAR4.GR1 cells and IFNAR1-reconstituted GAR4.AR1 cells have been described previously (Dunn et al., 2005). RMA-S is an MHC class I–deficient variant of the C57BL/6 strain T lymphoma RMA (Kärre et al., 1986). Tumor cells were propagated in vitro and injected s.c. in a volume of 150 µl endotoxin-free PBS into the shaved flanks of recipient mice as described previously (Dunn et al., 2005). Injected cells were >90% viable as assessed by trypan blue exclusion. Tumor size was measured on the indicated days and is presented as the mean of two perpendicular diameters. When calculating percent tumor growth, mice with tumors >6 mm in diameter were considered positive. Antibody treatment. For IFN-α/β receptor blockade, mice were injected i.p. with a single 2.5-mg dose of IFNAR1-specific MAR1-5A3 mAb (Sheehan et al., 2006) or GIR-208 isotype control mAb as described previously (Dunn et al., 2005). For IFN-γ neutralization, 750 µg of IFN-γ–specific H22 mAb (Schreiber et al., 1985) or PIP isotype control mAb was injected i.p. followed by a 250-µg dose every 7 d. Broad immunodepletion was achieved by i.p. administration of a mixture of anti-CD4 GK1.5 mAb (Dialynas et al., 1983), anti-CD8 YTS-169.4 mAb (Cobbold et al., 1984), and IFN-γ–specific H22 mAb. For this regimen, an initial dose of 750 µg of each mAb or of the control PIP mAb was followed by 250 µg of each every 7 d as described previously (Koebel et al., 2007). NK cell depletion was achieved in C57BL/6 mice by i.p. injection of 200 µg anti-NK1.1 PK136 mAb (Koo and Peppard, 1984; BioLegend) on days −2, 0, and 2 (relative to tumor injection) and 100 µg every 5 d. Generation of bone marrow chimeras. Recipient mice were irradiated with a single dose of 9.5 Gy and reconstituted with donor HSCs isolated from embryonic day (E) 14.5 fetal livers or 5-fluorouracil (5-FU)–treated adult bone marrow as described previously (Christensen et al., 2004; Dunn et al., 2005). For harvest of fetal liver cells (FLCs), embryos were extracted at E14.5, livers were removed, washed in sterile endotoxin-free PBS, and homogenized through a cell strainer using a syringe plunger. 5-FU–treated bone marrow was isolated 4–5 d after treatment of donor mice with 150 mg/kg 5-FU by i.p. injection. Cells were injected i.v. at a dose of 5 × 106 (FLCs) or 106 (5-FU–treated bone marrow) cells/mouse in 200 µl PBS. Total cell dose was determined by titration of FLCs (Fig. S3) or based on prior data (Dunn et al., 2005). For mixed chimeras, a 4:1 cell ratio was selected based on testing of different mixing ratios (Fig. S6). Animals were maintained on trimethoprim-sulfamethoxazole (Hi-Tech Pharmacal) antibiotic water prepared as described previously (Dunn et al., 2005) for 4 wk after irradiation, and tumor transplantation of chimeric mice was performed at least 12 wk after reconstitution. Hematopoietic reconstitution of all animals was verified by FACS staining of splenocytes at the completion of tumor transplantation experiments. Similar experimental results were obtained with mice reconstituted using FLCs or 5-FU–treated bone marrow as donor cells. Flow cytometry. Surface staining of single cell suspensions of splenocytes or tumor cells was performed using standard protocols and analyzed on a FACSCalibur (BD). Data analysis was conducted using FlowJo software (Tree Star). The following were obtained from BioLegend: anti-CD3-FITC (145-2C11), anti-CD4-PE (RMA4-5), anti-CD4-APC (GK1.5), anti–CD8α-APC (53-6.7), anti-CD8α-FITC (53-6.7), anti-B220-FITC (RA3-6B2), anti-CD11b-PE (M1/70), anti-CD11b-PerCP-Cy5.5 (and Pe-Cy7; M1/70), anti-DX5-PE (DX5), anti-DX5-APC (DX5), anti–Gr-1–FITC (RB6-8C5), anti-CD45-FITC (30-F11), anti-CD31-PE (MEC13.3), anti-CD24-FITC (M1/69), anti-CD103-PerCp-Cy5.5 (2E7), anti-Dec205-Pe-Cy7 (NLDC-145), anti-F4/80-PerCP-Cy5.5 (BM8), anti-CD11c-APC-Cy7 (N418), and SA-APC. Anti-CD11c-PE (HL3), anti-CD8α-PerCP-Cy5.5 (53–6.7), and anti-IFNGR1-biotin (GR20) were obtained from BD, anti-NKp46-PE (29A1.4) was purchased from eBioscience, and anti-IFNAR1-biotin (MAR1-5A3) was described previously (Sheehan et al., 2006). For pSTAT1 assays, splenocytes were stained for cell surface markers before stimulation with 10 ng/ml IFN-αv4 for 15 min. Cells were then fixed with 2% paraformaldehyde, permeabilized with 90% methanol, and stained for pSTAT1 (BD). For CD86 expression, cells were cultured for 18 h with 10 ng/ml IFN-αv4 before staining for cell surface markers and CD86-PE (BD). Tumor-specific CTL killing assay. Spleens were harvested from mice 20 d after tumor implantation, and single cell suspensions were prepared by homogenization using frosted glass slides. 4 × 107 splenocytes were cultured with 2 × 106 IFN-γ–treated, irradiated (100 Gy) tumor cells. 5 d later, the cells were harvested and used as CTL effector cells in a standard 4-h 51Cr-release cytotoxicity assay that used tumor cell targets seeded at 10,000 cells/well and pretreated with 100 U/ml IFN-γ for 48 h. For blocking assays, 10 µg/ml anti-CD8 (YTS-169.4), anti-CD4 (GK1.5), or control mAb (PIP) was added to the cell culture of effector and target cells. Percent specific killing was defined as (experimental condition cpm − spontaneous cpm)/(maximal (detergent) cpm − spontaneous cpm) × 100. NK cell cytotoxicity assay. Splenocytes were isolated from mice treated with 300 µg polyI:C (Sigma-Aldrich) by i.p. injection 24 h prior and were used as effector cells with 5,000 51Cr-labeled YAC-1 tumor targets. Percent specific killing was assessed after 4-h coincubation. Each sample was assayed in duplicate, and experiments were performed at least twice. Adoptive transfer of CD11c+ cells. Splenic CD11c+ cells from naive WT and Ifnar1−/− mice (10 mice/group) were positively selected by MACS (purity >90%) using CD11c microbeads (Miltenyi Biotec). 2 × 106 CD11c+ cells were mixed with 2 × 105 unedited MCA sarcoma cells (GAR4.GR1) in endotoxin-free PBS and injected s.c. in a volume of 200 µl into the flanks of Ifnar1−/− mice at day 0. 3 d later, 2 × 106 CD11c+ cells were injected s.c. around the site of tumor implantation. Antigen cross-presentation assay. DC cross-presentation of antigen to CD8+ OT-I T cells was assessed as previously described (Hildner et al., 2008). In brief, spleens from naive WT or Ifnar1−/− mice were digested with collagenase B (Roche) and DNase I (Sigma-Aldrich), and cellular subpopulations were isolated by MACS purification (Miltenyi Biotec). Total CD11c+ DCs were obtained by negative selection using B220, Thy1.2, and DX5 microbeads followed by positive selection with CD11c microbeads. CD8α+ DCs were recovered by B220, Thy1.2, DX5, and CD4 negative selection, followed by CD8α positive selection. CD4+ DCs were isolated by B220, Thy1.2, DX5, and CD8α negative selection, followed by CD4 positive selection. In all cases, purity of the population of interest was >97%. Splenocytes from Kb−/−Db−/−β2m−/− mice were prepared in serum-free medium, loaded with 10 mg/ml ovalbumin (EMD) by osmotic shock, and irradiated (13.5 Gy) as described previously (Hildner et al., 2008). OT-I T cells were purified from OT-I/Rag1−/− mice by CD11c and DX5 negative selection followed by positive selection with CD8α microbeads (purity >99%). T cells were fluorescently labeled by incubation with 1 µM CFSE (Sigma-Aldrich) for 9 min at 25°C at a density of 2 × 107 cells/ml. For the assay, 5 × 104 purified DCs were incubated with 5 × 104 CFSE-labeled OT-I T cells in the presence of varying numbers of irradiated, ovalbumin-loaded Kb−/−Db−/−β2m−/− splenocytes. In some assays, the irradiated target cells were mismatched (BALB/c) tumor cells expressing a truncated version of the IFN-γ receptor to render them IFN-γ insensitive and in which ovalbumin was retrovirally enforced (CMS-5-ΔIC). Ovalbumin expression was confirmed by coexpression of GFP by flow cytometry and by Western blot using a mouse antiovalbumin mAb (Santa Cruz Biotechnology, Inc.). After 3 d, cells were stained with anti-CD8α-APC and CFSE, or cell proliferation dye (eBioscience) dilution was measured by flow cytometry. For IFN-α treatment, recombinant mouse IFN-α5 (a gift from D. Fremont, Washington University in St. Louis) was added at 1,000 U/ml, whereas IFN-α/β receptor blockade was achieved by incubation with 5 µg/ml IFNAR1-specific MAR1-5A3 mAb. Online supplemental material. Fig. S1 shows the kinetics of tumor growth in mice treated with blocking IFNAR1-specific mAb. Fig. S2 demonstrates the importance of host IFN-γ sensitivity for rejection of unedited sarcomas. Fig. S3 presents a titration of FLCs for generation of bone marrow chimeras. Figs. S4 and S5 show the normal functional immune reconstitution of Ifngr1−/− bone marrow chimeras (Fig. S4) and the absence of radio-resistant, tissue-resident leukocytes in the tumors of these mice (Fig. S5). Fig. S6 shows a determination of the HSC mixing ratio used to generate mixed bone marrow chimeras. Fig. S7 shows an analysis of DC subsets in Ifnar1−/− mice. Fig. S8 shows further characterization of the Itgax-Cre+Ifnar1f/f mice. Fig. S9 shows adoptive transfer experiments of WT and Ifnar1−/− CD11c+ cells into Ifnar1−/− recipient mice. Fig. S10 shows decreased cross-presentation by CD8α+ DCs from Itgax-Cre+Ifnar1f/f mice compared with Ifnar1f/f mice using retrovirally transduced tumor cells as a source of antigen. Online supplemental material is available at http://www.jem.org/cgi/content/full/jem.20101158/DC1.

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Title: Journal of Clinical Investigation

Publisher: American Society for Clinical Investigation

ISSN (Print): 0021-9738

ISSN (Electronic): 1558-8238

Publication date Created: April 2 2018

Publication date (Electronic): March 5 2018

Publication date (Print): April 2 2018

Volume: 128

Issue: 4

Pages: 1413-1428

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DOI: 10.1172/JCI98047

PMC ID: 5873884

PubMed ID: 29504948

SO-VID: 706f77fc-0d22-4c33-91f1-79c04c3d5da0

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