Cancer Immunotherapy: Silencing Intracellular Negative Immune Regulators of Dendritic Cells

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Abstract

Dendritic cells (DCs) are capable of activating adaptive immune responses, or inducing immune suppression or tolerance. In the tumor microenvironment, the function of DCs is polarized into immune suppression that attenuates the effect of T cells, promoting differentiation of regulatory T cells and supporting tumor progression. Therefore, blocking negative immune regulators in DCs is considered a strategy of cancer immunotherapy. Antibodies can target molecules on the cell surface, but not intracellular molecules of DCs. The delivery of short-hairpin RNAs (shRNA) and small-interfering RNAs (siRNA) should be a strategy to silence specific intracellular targets in DCs. This review provides an overview of the known negative immune regulators of DCs. Moreover, a combination of shRNA/siRNA and DC vaccines, DNA vaccines in animal models, and clinical trials are also discussed.

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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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Inhibition of Allogeneic T Cell Proliferation by Indoleamine 2,3-Dioxygenase–expressing Dendritic Cells

Peter Terness, Thomas Bauer, Lars Röse … (2002)

Indoleamine 2,3-dioxygenase (IDO), an enzyme involved in the catabolism of tryptophan, is expressed in certain cells and tissues, particularly in antigen-presenting cells of lymphoid organs and in the placenta. It was shown that IDO prevents rejection of the fetus during pregnancy, probably by inhibiting alloreactive T cells, and it was suggested that IDO-expression in antigen-presenting cells may control autoreactive immune responses. Degradation of tryptophan, an essential amino acid required for cell proliferation, was reported to be the mechanism of IDO-induced T cell suppression. Because we wanted to study the action of IDO-expressing dendritic cells (DCs) on allogeneic T cells, the human IDO gene was inserted into an adenoviral vector and expressed in DCs. Transgenic DCs decreased the concentration of tryptophan, increased the concentration of kynurenine, the main tryptophan metabolite, and suppressed allogeneic T cell proliferation in vitro. Kynurenine, 3-hydroxykynurenine, and 3-hydroxyanthranilic acid, but no other IDO-induced tryptophan metabolites, suppressed the T cell response, the suppressive effects being additive. T cells, once stopped in their proliferation, could not be restimulated. Inhibition of proliferation was likely due to T cell death because suppressive tryptophan catabolites exerted a cytotoxic action on CD3+ cells. This action preferentially affected activated T cells and increased gradually with exposure time. In addition to T cells, B and natural killer (NK) cells were also killed, whereas DCs were not affected. Our findings shed light on suppressive mechanisms mediated by DCs and provide an explanation for important biological processes in which IDO activity apparently is increased, such as protection of the fetus from rejection during pregnancy and possibly T cell death in HIV-infected patients.

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DNA Vaccines—How Far From Clinical Use?

Dominika Hobernik, Matthias Bros (2018)

Two decades ago successful transfection of antigen presenting cells (APC) in vivo was demonstrated which resulted in the induction of primary adaptive immune responses. Due to the good biocompatibility of plasmid DNA, their cost-efficient production and long shelf life, many researchers aimed to develop DNA vaccine-based immunotherapeutic strategies for treatment of infections and cancer, but also autoimmune diseases and allergies. This review aims to summarize our current knowledge on the course of action of DNA vaccines, and which factors are responsible for the poor immunogenicity in human so far. Important optimization steps that improve DNA transfection efficiency comprise the introduction of DNA-complexing nano-carriers aimed to prevent extracellular DNA degradation, enabling APC targeting, and enhanced endo/lysosomal escape of DNA. Attachment of virus-derived nuclear localization sequences facilitates nuclear entry of DNA. Improvements in DNA vaccine design include the use of APC-specific promotors for transcriptional targeting, the arrangement of multiple antigen sequences, the co-delivery of molecular adjuvants to prevent tolerance induction, and strategies to circumvent potential inhibitory effects of the vector backbone. Successful clinical use of DNA vaccines may require combined employment of all of these parameters, and combination treatment with additional drugs.

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Author and article information

Journal

Journal ID (nlm-ta): Cancers (Basel)

Journal ID (iso-abbrev): Cancers (Basel)

Journal ID (publisher-id): cancers

Title: Cancers

Publisher: MDPI

ISSN (Electronic): 2072-6694

Publication date (Electronic): 17 January 2019

Publication date Collection: January 2019

Volume: 11

Issue: 1

Electronic Location Identifier: 108

Affiliations

[1 ]Department of Emergency Medicine, Kaohsiung Medical University Hospital, Kaohsiung Medical University, Kaohsiung 807, Taiwan; 980542@ 123456ms.kmuh.org.tw (Y.-H.L.); 910201@ 123456ms.kmuh.org.tw (I.-J.Y.); 890077@ 123456ms.kmuh.org.tw (K.-T.L.)

[2 ]Department of Biochemistry and Molecular Biology, College of Medicine, National Cheng Kung University, Tainan 701, Taiwan; a1211207@ 123456mail.ncku.edu.tw

[3 ]Institute of Basic Medical Sciences, College of Medicine, National Cheng Kung University, Tainan 701, Taiwan

[4 ]School of Medicine, College of Medicine, Kaohsiung Medical University, Kaohsiung 807, Taiwan

[5 ]Graduate Institute of Clinical Medicine, College of Medicine, Kaohsiung Medical University, Kaohsiung 807, Taiwan; kuopolin@ 123456seed.net.tw

Author notes

[* ]Correspondence: yohoco@ 123456gmail.com

[†]

Contributed equally.

Author information

Po-Lin Kuo https://orcid.org/0000-0003-2487-2818

Article

Publisher ID: cancers-11-00108

DOI: 10.3390/cancers11010108

PMC ID: 6357062

PubMed ID: 30658461

SO-VID: 5c9218d3-4e8f-4b00-ada8-957a37f817b8

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Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license ( http://creativecommons.org/licenses/by/4.0/).

Cancer Immunotherapy: Silencing Intracellular Negative Immune Regulators of Dendritic Cells

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