Exogenous IL-33 Restores Dendritic Cell Activation and Maturation in Established Cancer

Donye Dominguez, Cong Ye, Zhe Geng, Siqi Chen, Jie Fan, Lei Qin, Alan Long, Long Wang, Zhuoli Zhang, Yi Zhang, Deyu Fang, Timothy M. Kuzel and Bin Zhang
J Immunol February 1, 2017, 198 (3) 1365-1375; DOI: https://doi.org/10.4049/jimmunol.1501399

Abstract

The role of IL-33, particularly in tumor growth and tumor immunity, remains ill-defined. We show that exogenous IL-33 can induce robust antitumor effect through a CD8+ T cell–dependent mechanism. Systemic administration of rIL-33 alone was sufficient to inhibit growth of established tumors in transplant and de novo melanoma tumorigenesis models. Notably, in addition to a direct action on CD8+ T cell expansion and IFN-γ production, rIL-33 therapy activated myeloid dendritic cells (mDCs) in tumor-bearing mice, restored antitumor T cell activity, and increased Ag cross-presentation within the tumor microenvironment. Furthermore, combination therapy consisting of rIL-33 and agonistic anti-CD40 Abs demonstrated synergistic antitumor activity. Specifically, MyD88, an essential component of the IL-33 signaling pathway, was required for the IL-33–mediated increase in mDC number and upregulation in expression of costimulatory molecules. Importantly, we identified that the IL-33 receptor ST2, MyD88, and STAT1 cooperate to induce costimulatory molecule expression on mDCs in response to rIL-33. Thus, our study revealed a novel IL-33–ST2–MyD88–STAT1 axis that restores mDC activation and maturation in established cancer and, thereby, the magnitude of antitumor immune responses, suggesting a potential use of rIL-33 as a new immunotherapy option to treat established cancer.

Introduction

The new IL-1 family member, IL-33 (1), is mainly and constitutively expressed by nonhematopoietic cells, such as fibroblasts, epithelial cells, and endothelial cells (2). IL-33 was identified as the functional ligand for the orphan receptor ST2 (IL-1R–like-1) (3, 4). IL-33 signaling via ST2 and IL-1R accessory protein dimers (5) results in the recruitment of MyD88, which induces activation of various signaling pathways, including NF-κB and MAPK pathways (1, 4, 6–8). Accumulating evidence demonstrates that IL-33 has an important role in promoting allergic responses (4, 9) and other Th2-related diseases, such as asthma (10), atopic dermatitis (11), and anaphylaxis (12). In contrast, IL-33 may also protect against inflammation-associated atherosclerosis (13) or infection-induced tissue damage (14, 15). Therefore, IL-33 has a dual functionality in different diseases, depending on the immune mechanism underlying the pathogenesis of various disease conditions.

Although IL-33 has been studied primarily for its role in the context of inflammation, allergy, and autoimmunity (1, 6, 16), its role in tumor immunity and tumor growth is only beginning to be appreciated (17). It is now becoming clear that IL-33 plays a role far beyond the realm of Th2 immunity by promoting Th1 immune responses (18), regulating the development of antiviral CD8+ T cells, and driving the effector function of CD8+ T cells (19, 20). There are several recent studies showing that IL-33 can promote antitumor CD8+ T cell responses in experimental mouse tumor models (21–23). However, the molecular and cellular mechanisms of how IL-33 influences the antitumor CD8+ T cell responses remain elusive.

Tumor can escape host immune surveillance by fostering a highly suppressive microenvironment (24–26). There is clear evidence for the role of tumor-induced dendritic cell (DC) dysfunction as one important mechanism for tumor-induced immune escape (27, 28), because DCs function as key professional APCs to induce tumor-specific immune responses, particularly via cross-priming through MHC class I Ag presentation (29, 30). DC defects are caused by abnormal differentiation leading to decreased production of fully competent APCs and increased accumulation of immature tolerogenic DCs. In fact, the inability to mount a potent antitumor immune response has often been attributed to DC defects. Limited data are available on the role of IL-33 in DC activity. IL-33 was shown to promote bone marrow DC generation in vitro (31). Moreover, IL-33–activated DCs drove an atypical Th2-type immune response (32) and exacerbated allergic lung inflammation (33). However, no study has examined the effects of IL-33 on DC phenotype and function in the context of cancer.

In this study, we show that systemic administration of rIL-33 alone was sufficient to inhibit the growth of established B16 melanoma, EG7 lymphoma, and clinically more relevant BRAFV600EPTEN inducible melanomas. Furthermore, the therapeutic efficacy of rIL-33 was primarily dependent on CD8+ T cells rather than NK+ cells and CD4+ T cells. In addition to CD8+ T cells, dysfunctional myeloid DCs (mDCs) of tumor-bearing mice were activated directly by rIL-33 to express costimulatory molecules potentiating antitumor immune responses. As a result, synergistic antitumor activity was achieved with rIL-33 and agonistic anti-CD40 combination therapy. Mechanistically, we identified a novel IL-33–ST2–MyD88–STAT1 axis that regulated DC activation and maturation and, thereby, the magnitude of antitumor immune responses. Thus, our results may have direct impact for developing rIL-33 as a novel and promising approach to treat established cancer.

Materials and Methods

Mice, cell lines, and reagents

C57BL/6 wild-type (WT), C57BL/6 MyD88−/−, Rag1−/−, BRAFV600EPTEN (Tyr::CreER; BrafCA/+; Ptenlox5/lox5), and Pmel-1 mice were purchased from the Jackson Laboratory. OT-1 Rag1−/− mice were purchased from Taconic. Dr. H. Schreiber (University of Chicago) provided EG7, B16F10, B16-OVA, and B16-SIY cell lines, SIYRYYGL (SIY) peptides, and 2C-transgenic mice. The BPS-1 mouse melanoma was generated from a spontaneously arising tumor in BRAFV600EPTEN (Tyr::CreER; BrafCA/+; Ptenlox5/lox5)-transgenic mice (34, 35). We demonstrated the presence of the BRAFV600E transversion in BPS-1 cells, and Braf inhibitor PLX4032 can effectively inhibit the growth of BPS-1 melanomas in vivo (data not shown). All of the cell lines were routinely tested for mycoplasma infections by culture and DNA stain and were maintained in complete medium composed of RPMI 1640 with 5% FBS. All animal experiments were approved by institutional animal use committees of the University of Texas Health Science Center at San Antonio and Northwestern University. H-2Db/gp100 tetramers were provided by the National Institutes of Health Tetramer Core Facility. 1B2 Abs (clonotypic anti-2C TCR) were provided by Dr. J. Kline (University of Chicago). Phospho-STAT mAbs were purchased from Cell Signaling Technology. All other flow cytometry mAbs were obtained from eBioscience and BioLegend. Proliferation dye eFluor 450 was from eBioscience. 4-hydroxytamoxifen (4-HT; Z-isomer) was purchased from Sigma-Aldrich. Agonistic anti-CD40 (clone FGK4.5), depleting mAb clone GK1.5 (anti-CD4), clone 53.6.7 (anti-CD8α), clone PK136 (anti-NK1.1), and control IgG were purchased from Bio X Cell. Recombinant murine IL-33 and GM-CSF were purchased from BioLegend. ST2-blocking Ab, clone DJ8, was purchased from MD Bioproducts. The peptide hgp10025–33 was purchased from GenScript. For MTT assay, a Cell Proliferation Kit I (MTT) was purchased from Roche.

Analysis of cells by flow cytometry

All samples were initially incubated with 2.4G2 to block Ab binding to Fc receptors. Single-cell suspensions were stained with 1 μg of relevant mAbs and then washed twice with cold PBS. H-2Db/gp100 tetramer staining, Foxp3 staining, and intracellular IFN-γ staining were performed as previously described (36). For STAT staining, cells were treated as indicated and then 106 cells were fixed in 4% formaldehyde for 10 min at room temperature. Cells were washed with ice-cold PBS containing 2% BSA, followed by another wash step with ice-cold PBS. Cells were resuspended in 80% methanol and incubated for 30 min at −20°C. The pellet was washed twice with ice-cold PBS. mAbs were added in a final volume of 100 μl of ice-cold PBS and incubated at 4°C for 45 min. Cells were washed with ice-cold PBS containing 2% BSA and analyzed by flow cytometry. Samples were processed using a MACSQuant Analyzer (Miltenyi Biotec), and data were analyzed with FlowJo software.

T cell and DC purification

Splenic CD8+ T cells from WT, 2C, Pmel-1, and OT-1 mice were selected using IMag CD8 magnetic particles (BD Biosciences) with >95% purity routinely checked by flow cytometry. For DC selection, draining lymph nodes (DLNs) or spleen cells were suspended in cold PBS at a concentration of 108 cells per milliliter and incubated with anti-mouse CD11c-Biotin for 30 min at 4°C. Cells were then washed and incubated with magnetic streptavidin particles for 30 min (Streptavidin Particles Plus-DM; BD Biosciences) and purified according to the manufacturer’s protocol with an expected purity of 80–90%.

ELISPOT assay

For evaluation of ex vivo DC priming ability, an ELISPOT assay was performed. Ninety-six well MultiScreen-IP Filter Plates (Millipore) were used for plating; all other reagents were from the Mouse IFN gamma ELISPOT Ready-SET-Go (eBioscience). All steps were done according to the manufacturer’s protocol. Each well contained 105 DCs along with 105 CD8+ T cells in a final volume of 200 μl per well. Negative controls were CD8+ T cells alone. Samples were incubated for 48 h before harvest. The numbers and diameters of spots were counted in triplicates and calculated by an automatic ELISPOT counter.

In vitro cell treatment

For in vitro treatment of DCs purified from tumor-bearing mice or tumor-free mice, rIL-33 was added at 10 ng/ml for 10 min before STAT staining. To exclude the potential role of T cells in IL-33–induced DC activation, T cells were depleted from splenocytes using CD90.1 selection prior to purification of CD3−CD11c+ cells by FACS for >99% purity of cells. Sorted CD3−CD11c+ WT DCs and eFluor 450–labeled MyD88−/− DCs were mixed at a 1:1 ratio and treated or not with rIL-33. The next day, cells were analyzed for STAT1 phosphorylation by flow cytometry. WT DCs were gated as e450−, whereas MyD88−/− DCs were CD3−CD11c+e450+. Anti-ST2 (clone DJ8) was added at 10 μg/ml when used. For in vitro treatment of CD8+ T cells, purified splenic CD8+ T cells from tumor-free mice were labeled with eFluor 450, as described previously (36). rIL-33 was added at 10 ng/ml for 48 h before proliferation and IFN-γ analysis.

Tumor challenge and treatments

B16-F10, B16-SIY, BPS-1, or EG7 cells (1 × 106) in suspension were injected s.c. into the rear right flank of mice. For the DC-transfer experiment, purified splenic CD11c+ DCs from WT or MyD88−/− mice were stimulated with rIL-33 (10 ng/ml). Three days later, B16-SIY–bearing mice were injected with these IL-33–activated WT or MyD88−/− DCs. The following day, naive 2C CD8+ T cells were stained with Cell Proliferation Dye eFluor 450 and transferred i.v. After 4 d, DLNs and tumor tissues were harvested to measure proliferation and IFN-γ production of transferred 2C T cells. For in vivo rIL-33 treatment, groups of mice received daily i.p. injection of PBS or 1 μg of rIL-33 dissolved in PBS, starting on day 8–10 after injection or when tumors reached ∼150–300 mm3. Depletion of CD4+ T cells, CD8+ T cells, or NK cells was achieved as previously described (36) by twice-a-week i.p. injection of depleting mAb clone GK1.5 (anti-CD4; 200 μg), clone 53.6.7 (anti-CD8α; 200 μg), or clone PK136 (anti-NK1.1; 200 μg) starting 1 d prior to rIL-33 treatment. Induction of melanoma in BRAFV600EPTEN mice was performed as described previously (BRAF cooperates with PTEN loss to induce metastatic melanoma) (34, 35). 4-HT was dissolved to 2 mg/ml in DMSO, and 4–6 μl was painted on the hind flank for three consecutive days. For the BRAFV600EPTEN model, rIL-33 treatment was initiated when tumors became measureable. For combination therapy, agonistic anti-CD40 (clone FGK4.5; 150 μg) was administered i.p. at the same time or 1 d after initiation of rIL-33 treatment and was subsequently administered every third day. In all experiments, tumor size was determined at 2–3-d intervals. Tumor volumes were measured along orthogonal axes (a, b, and c) and calculated as abc/2.

Statistical analysis

Mean values were compared using an unpaired Student two-tailed t test. The statistical differences between the survival of groups of mice were calculated using the log-rank test. Probability values > 0.05 were considered nonsignificant.

Results

Antitumor effect of rIL-33 is dependent on CD8+ T cells

The effects of exogenous IL-33 on tumorigenesis were reported in several recent studies (21–23). However, the exact roles of IL-33 in regulating antitumor immunity and antitumor growth are barely understood. We examined the effect of daily i.p. injection of rIL-33 on the growth of B16F10 melanoma. Given the importance of tumor size and duration of tumor growth in experimental therapy for clinical implication, mice bearing established tumors (∼150–250 mm3) were randomized and treated beginning on day 8–10. A dose-response relationship for the rIL-33–mediated effect on tumor growth (Fig. 1A) and mice survival (Fig. 1B) was initially demonstrated from 0.5 to 1.0 μg i.p. daily. Similarly, systemic administration of rIL-33 (1 μg/d) significantly inhibited the growth of established EG7 lymphoma (Fig. 1C). The in vivo antitumor activity of rIL-33 was dependent on adaptive antitumor immune responses, because rIL-33 therapy was ineffective against B16F10 tumors in Rag1−/− mice (lacking T and B lymphocytes) (Fig. 1D). As expected, rIL-33 did not affect B16F10 tumor growth in vitro (data not shown), consistent with previous studies (23). Furthermore, depletion of CD8α+ cells, rather than CD4+ cells or NK cells, abrogated the tumor-inhibiting activity of rIL-33 treatment in EG7-bearing mice (Fig. 1E) and B16F10-bearing mice (Fig. 1F), which indicates that the tumor-inhibiting activity of rIL-33 was primarily dependent on CD8+ cells and independent of CD4+ cells or NK cells.

FIGURE 1.
FIGURE 1.

Administration of rIL-33 alone is sufficient to inhibit tumor growth in a CD8+ T cell–dependent manner. (A and B) The rIL-33–induced antitumor effect is dose dependent. C57BL/6 mice were injected s.c. with 106 B16-F10 cells and treated daily with the indicated dose of rIL-33 (i.p.) starting from day 9. In (B), mice were sacrificed when tumor volume reached 2 cm3. *p < 0.05, **p < 0.01, PBS versus 0.5 or 1 μg/d treatment. (C) Inhibition of EG7 tumor growth by rIL-33. C57BL/6 mice were injected s.c. with 106 EG7 cells and treated daily with 1 μg rIL-33 (i.p.) starting from day 11. **p < 0.01. (D) Tumor inhibition by rIL-33 is dependent on adaptive immunity. C57BL/6 Rag1−/− mice were injected s.c. with 106 B16-F10 cells and treated daily with 1 μg rIL-33 (i.p.) starting from day 10. (E and F) rIL-33 inhibits tumor growth in a CD8+ T cell–dependent manner. Depletion of CD4+ T cells, CD8+ T cells, or NK cells was achieved by twice-weekly injection of anti-CD4, anti-CD8, or anti-NK1.1 depleting Abs, respectively, on the same day in EG7-bearing mice (E) or 2 d prior to rIL-33 or PBS administration in B16-F10–bearing mice (F). In all experiments, rIL-33 was administered when tumor volume was ∼150–250 mm3. Data (mean ± SEM) are representative of at least three independent experiments with five mice per group. *p < 0.05.

rIL-33 treatment promotes antitumor T cell immunity

We next characterized exogenous rIL-33–induced antitumor T cell immunity. Phenotype and cytokine profiles of tumor-infiltrating immune cells were examined 8–10 d after rIL-33 or PBS treatment. We found that rIL-33 treatment remarkably enhanced the infiltration of CD4+, CD8+ T cells (Fig. 2A, 2B) and tumor Ag gp100-specific CD8+ T cells (Fig. 2C). Moreover, there was significantly increased IFN-γ production (Fig. 2D) and KLRG1 expression (data not shown) by tumor-infiltrating CD8+ T cells from rIL-33–treated mice compared with the PBS group. The percentage of Foxp3+ cells among CD4+ T cells remained unchanged. There was a trend toward an increase in the absolute number of CD4+Foxp3+ regulatory T cells (Tregs) by rIL-33 treatment, but the difference did not reach statistical significance (Fig. 2E). rIL-33 treatment resulted in a significant increase in the absolute number of tumor-infiltrating Gr1+CD11b+ myeloid-derived suppressor cells (MDSCs) (Fig. 2B). In addition, IL-33 was effective in inhibiting the growth of immunogenic B16-SIY (expressing SIY Ag) (Supplemental Fig. 1A), which also corresponded to the infiltration of greater numbers of CD8+ T cells but fewer CD4+Foxp3+ Tregs (Supplemental Fig. 1B, 1C). IL-33 treatment promoted CD8+ effector T cell proliferation within B16-SIY tumors, as measured by the expression of the cell cycle–associated protein Ki67 (Supplemental Fig. 1D), as previously reported. In B16-F10 (Fig. 2F) and B16-SIY (Supplemental Fig. 1E) tumor models, intratumoral ratios of CD8+ T effector cells/Tregs and CD8+ T effector cells/MDSCs were markedly increased by rIL-33 therapy. Furthermore, we observed a significant increase in splenic frequencies of Tregs (Supplemental Fig. 2A) and MDSCs (Supplemental Fig. 2B) in IL-33–treated mice. Collectively, these data suggest that systemic administration of rIL-33 promoted Ag-specific CD8+ T cell expansion and effector function, despite the fact that this expansion was accompanied by an expansion of Tregs and MDSCs in the periphery and tumor.

FIGURE 2.
FIGURE 2.

rIL-33 therapy increases antitumor CD8+ T cell immunity. Percentage (A) and absolute number (B) of B16-F10 tumor-infiltrating TCRVb+CD4+, TCRVb+CD8+, and Gr1+CD11b+ cells, as determined by flow cytometry (n = 5). (C) rIL-33 treatment increased gp100-specific tetramer+TCRVb+CD8+ cells per 106 tumor infiltrates, as calculated by flow cytmometry (n = 10). (D) rIL-33 treatment increased IFN-γ production from tumor-infiltrating CD8+ cells, as determined by flow cytometry (n = 10). (E) Percentage and absolute number of B16-F10 tumor-infiltrating CD4+Foxp3+ cells, as determined by flow cytometry (n = 5). (F) IL-33 shifts the ratio of CD8+ T cells/suppressive cells in favor of an increased proportion of CD8+ T cells, as determined by flow cytometry percentages. Cells were collected for flow cytometry 8–10 d after PBS or rIL-33 treatment. Data (mean ± SEM) are representative of at least two independent experiments. *p < 0.05, **p < 0.01.

Defective tumor-associated DC function is reversed by exogenous IL-33

Although we showed that rIL-33 had a direct effect on CD8+ T cell expansion and IFN-γ production (Supplemental Fig. 3), it is very likely that IL-33 can potentiate antitumor T cell immunity through other immune cells, such as DCs. Indeed, in B16-SIY–bearing mice, the absolute number of tumor-infiltrating CD11b+CD11c+ mDCs was greatly increased by rIL-33 treatment (Fig. 3A). IL-33 also induced upregulation of CD40 among these DC infiltrates (Fig. 3B). Similarly, there was significantly increased expression of CD40 and CD80 in the IL-33–treated B16-F10–bearing mice compared with the control group (data not shown). To further examine the functional significance of these tumor-associated DCs during rIL-33 therapy, CD11c+ DCs were purified from PBS-treated or rIL-33–treated B16-SIY–bearing mice. As shown in Fig. 3C, DCs from IL-33–treated mice induced more IFN-γ–producing SIY-specific 2C CD8+ T cells, indicating the possibility that more DCs from IL-33–treated mice than from PBS-treated mice present the SIY Ag, resulting in more 2C T cells recognizing their cognate peptides. Likewise, in B16-OVA–bearing mice, DCs from IL-33–treated mice, but not from PBS-treated mice, were able to cross-present OVA Ag to trigger IFN-γ production from OT-1 CD8+ T cells (Supplemental Fig. 4). The data indicate that exogenous IL-33 may induce DC activation and maturation within the tumor microenvironment, thereby promoting antitumor T cell immunity.

FIGURE 3.
FIGURE 3.

IL-33 activates tumor-associated DCs to restore their T cell cross-priming ability. (A) rIL-33 increased the absolute number of B16-SIY–infiltrating DCs gated as CD45+Gr1−CD11b+CD11c+. (B) Percentage and median fluorescence intensity (MFI) of CD40, CD80, CD86, and MHC-II expression were determined among tumor-infiltrating DCs (n = 10). (C) DCs from IL-33–treated B16-SIY–bearing mice have rescued cross-presentation and priming ability compared with those from PBS-treated B16-SIY–bearing mice, as quantified with an IFN-γ–based ELISPOT assay. DCs were cultured with naive 2C CD8+ T cells without the addition of exogenous Ag at a 1:1 ratio for 48 h. Representative ELISPOT panel is shown on the left (n = 3). Data (mean ± SEM) are representative of two independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001.

Antitumor activity of exogenous IL-33 against de novo BRAFV600EPTEN-driven melanoma

Given the potential limitation of the transplantable tumor models above, we further examined the antitumor effect of IL-33 administration in a more clinically relevant de novo melanoma model (i.e., the BRAFV600EPTEN mouse model) (34, 35). As shown in Fig. 4A, systemic administration of rIL-33 inhibited the growth of established BRAFV600EPTEN-inducible melanomas. Moreover, we observed significantly increased numbers of CD8+ T cells producing IFN-γ in rIL-33–treated mice compared with the control group (Fig. 4B). rIL-33 treatment also augmented expression levels of CD80 and CD86, but not CD40 or MHC class II (MHC-II), on mDCs from DLNs (Fig. 4C). Furthermore, DCs from rIL-33–treated mice had increased ability to cross-prime Ag-specific CD8+ T cells (Fig. 4D). Thus, the data obtained from the BRAFV600EPTEN melanoma model are in line with those from the transplantable tumor models above, suggesting that rIL-33 treatment directly activates the tumor-associated mDCs in vivo to express costimulatory surface molecules, thereby boosting antitumor T cell immunity. The discrepancy of IL-33–activated costimulatory molecules between B16 and BRAFV600EPTEN models may be explained by the different nature of each tumor model tested and the time of harvest.

FIGURE 4.
FIGURE 4.

rIL-33 inhibits tumor growth and promotes the T cell cross-priming ability of tumor-associated DCs in the BRAFV600EPTEN model. (A) rIL-33 inhibited melanoma growth in the BRAFV600EPTEN model. Forty-two days after 4-HT induction, mice were treated with IL-33 (1 μg/d) or PBS for 25 d. Fold change is between the start and the end of treatment. (B) rIL-33 treatment increased IFN-γ production from CD8+ cells from PBS- or IL-33–treated melanoma-bearing mice, as determined by flow cytometry (n = 3). (C) Percentage and median fluorescence intensity (MFI) of CD40, CD80, CD86, and MHC-II expression were determined among DLN CD11b+CD11c+Gr-1− DCs (n = 3). (D) DCs from IL-33–treated melanoma-bearing mice have increased T cell cross-priming ability compared with those from PBS-treated melanoma-bearing mice, as quantified with an IFN-γ–based ELISPOT assay. Purified CD11c+ cells from DLNs were cultured with naive Pmel CD8+ T cells, without the addition of exogenous Ag, at a 1:1 ratio for 48 h. Representative ELISPOT panel is shown on the left (n = 3). Data (mean ± SEM) are representative of two independent experiments. *p < 0.05, **p < 0.01.

Synergistic antitumor activity of rIL-33 and agonistic anti-CD40 combination therapy

rIL-33 significantly upregulated CD40 expression in tumor-associated DCs in vivo (Fig. 3B) and promoted DC-mediated antitumor T cell activity (Fig. 3C). Therefore, it is reasonable to combine agonistic anti-CD40 Ab therapy and IL-33 administration in cancer treatment. Consistent with published data (37), we confirmed the significant antitumor activity of anti-CD40 alone and the comparable effect of rIL-33 alone against established B16-F10 (Fig. 5A) and B16-SIY (Fig. 5B) tumors. More strikingly, the antitumor activity of anti-CD40 was significantly enhanced by rIL-33 treatment, with all tumor-bearing mice efficiently arresting and inducing the regression of some established B16-F10 (Fig. 5A) and B16-SIY (Fig. 5B) tumors. A similar combinatorial synergy was observed for IL-33 and anti-CD40 mAb, but it had a lower efficacy in suppressing the growth of the more immune-resistant BPS-1 melanomas (Fig. 5C) that originated from a spontaneously arising tumor in BRAFV600EPTEN mice. The reduced efficacy of anti-CD40/rIL-33 against BPS-1 melanomas correlated with a lower frequency of CD8+ T cells naturally infiltrating the tumors (1.5 ± 1.4%) compared with B16 tumors (Fig. 2B). As expected, rIL-33 or anti-CD40 alone increased tumor infiltration of CD8+ T cells and IFN-γ production from these CD8+ T cells. Combination rIL-33 and anti-CD40 therapy showed further improved activity over either treatment alone (Fig. 5D).

FIGURE 5.
FIGURE 5.

Combination of anti-CD40 and rIL-33 has a synergistic effect to delay tumor growth. C57BL/6 mice (n = 5) were injected s.c. with 106 B16-F10 (A) or B16-SIY (B) tumor cells and received PBS or rIL-33 (1 μg/d) daily starting from day 8. Agonistic anti-CD40 was administered i.p. every 3 d starting from day 9. (C) Inhibition of BPS-1 tumor growth by combination therapy. Mice (n = 5) were injected s.c. with 106 BPS-1 tumor cells and received the same therapy as in (A) and (B). (D) rIL-33 treatment increased the number of B16-SIY–infiltrating CD8+ and CD8+ IFN-γ+ T cells. Data (mean ± SEM) are representative of two independent experiments. *p < 0.05, ***p < 0.001.

Exogenous IL-33–mediated antitumor effect requires MyD88

IL-33 signals through the adaptor protein MyD88 (38); however, the role of MyD88 in regulating tumor immunity by IL-33 has not been studied in detail. We found that rIL-33 treatment delayed the growth of established B16-SIY tumors in WT mice, but this antitumor effect was completely abrogated in MyD88−/− mice (Fig. 6A), suggesting an essential role for MyD88 in rIL-33–mediated antitumor activity. Furthermore, rIL-33 treatment failed to enhance the frequency, absolute number, and proliferation (Ki-67 expression) of intratumoral CD8+ T cells from MyD88−/− mice compared with WT mice (Fig. 6B), as would be predicted. We also noticed that MyD88 signaling was dispensable for tumor growth and antitumor CD8+ T cell activity in the absence of rIL-33 therapy (Fig. 6).

FIGURE 6.
FIGURE 6.

rIL-33–mediated antitumor effect is dependent on the MyD88 signaling pathway. (A) C57BL/6 WT or MyD88−/− mice were injected s.c. with 106 B16-SIY tumor cells and received PBS or rIL-33 (1 μg/d) daily starting from day 9 (n = 5). (B) Percentage and absolute number of tumor-infiltrating TCRVb+CD8+ T cells and percentage of Ki-67+ in tumor-infiltrating CD8+ T cells, as determined by flow cytometry (n = 5). MyD88−/− mice exhibited decreased CD8+ T cell infiltration and proliferation in response to rIL-33. Data (mean ± SEM) are representative of two independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001.

Exogenous IL-33 signaling in DCs requires ST2, MyD88, and STAT1

Given the ability of rIL-33 to rescue tumor-associated DC activity, we tested whether rIL-33 therapy acts on tumor-associated DCs through MyD88 signaling. Examination of tumor-free MyD88−/− mice showed similar proportions of mDCs in the lymph nodes and spleen compared with WT mice (data not shown), indicating that MyD88 is not required for DC development or differentiation. However, in the B16-SIY model, IL-33 treatment increased the absolute number of intratumoral DCs in WT mice rather than in MyD88−/− mice (Fig. 7A). We also observed an upregulation of costimulatory molecules (CD86, CD40, CD80, and MHC-II) on DCs from DLNs in WT tumor-bearing mice in response to rIL-33 therapy, but this effect was completely abrogated in MyD88−/− mice (Fig. 7B). To further dissect the molecular mechanism of MyD88-mediated signaling downstream of exogenous IL-33 in DC activation, we examined the involvement of STAT activation by IL-33. As shown in Fig. 7C, rIL-33 treatment specifically induced phosphorylation of STAT1, but not STAT5, in mDCs from WT tumor-bearing mice (data not shown). More interestingly, the frequency of ST2+STAT1+ cells in mDCs was reduced significantly in response to rIL-33 therapy in tumor-bearing MyD88−/− mice compared with WT mice (Fig. 7D), suggesting a potential functional link between STAT1 and ST2-MyD88–mediated signaling downstream of IL-33.

FIGURE 7.
FIGURE 7.

IL-33 signaling in tumor-associated DCs requires MyD88 and STAT1. (A) rIL-33 increased the number of B16-SIY–infiltrating Gr-1−CD11b+CD11c+ DCs from WT mice but not from MyD88−/− mice (n = 5). (B) Representative flow cytometry analysis of the frequency of CD40, CD80, CD86, or MHC-II in DCs from DLNs of WT or MyD88−/− tumor-bearing mice (n = 5) killed 8–10 d after PBS or rIL-33 treatment. Internal controls were gated from cell populations within the analyzed sample that do not express DC costimulatory molecules. (C) Gr1−CD11c+ DCs from DLNs of tumor-bearing WT mice were rested overnight and then stained intracellularly for phosphorylated STAT1 by flow cytometry. (D) Representative flow cytometry analysis of phosphorylated STAT1 and ST2 by gated CD11b+CD11c+ DCs from DLNs of rIL-33–treated tumor-bearing WT or MyD88−/− mice. Data (mean ± SEM) are representative of three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001.

We next examined whether MyD88 signaling in the DC compartment was responsible for IL-33–mediated CD8+ T cell priming. As expected, WT DCs, but not MyD88−/− DCs, increased costimulatory molecules and STAT1 in response to rIL-33 (Fig. 8A). Furthermore, transfer of rIL-33–activated WT DCs triggered a greater proliferation of Ag-specific CD8+ T cells in DLNs from B16-SIY–bearing mice than did transfer of rIL-33–activated MyD88−/− DCs (Fig. 8B). Similarly, a significant increase in the frequencies of tumor-infiltrating CD8+IFN-γ+ T cells and proliferating Ki-67+CD8+ T cells was observed in these B16-SIY–bearing mice following the transfer of IL-33–activated WT DCs compared with MyD88−/− DCs (Fig. 8C).

FIGURE 8.
FIGURE 8.

IL-33 induces DC maturation and T cell priming via MyD88. (A) rIL-33 increased costimulatory molecules and STAT1 on WT DCs but not on MyD88−/− DCs. Purified splenic DCs were stimulated with 10 ng/ml of IL-33 daily for 3 d. (B) IL-33–activated DCs in (A) were injected into mice harboring established B16-SIY tumors (n = 5). The following day, naive 2C CD8+ T cells (1B2+) were stained with Cell Proliferation Dye eFluor 450 and transferred i.v. After 4 d, DLNs were harvested to measure proliferation of transferred cells. FlowJo software was used to calculate the percentage of individual generations by eFluor 450 dye dilution. (C) Flow cytometry was used to analyze the percentages of IFN-γ+ or Ki-67+ cells among 2C CD8+ T cells in tumor tissues. Data (mean ± SEM) are representative of at least two independent experiments. *p < 0.05, **p < 0.01.

To test whether ST2 signal could promote DC activation and maturation, we activated WT splenic DCs from tumor-bearing mice in the presence of GM-CSF and rIL-33, with or without anti-ST2–blocking mAb. Consistent with a requirement for MyD88 for IL-33–induced DC activation, as shown in Fig. 9A, expression of costimulatory molecules (CD86, CD40, CD80, and MHC-II) was upregulated by rIL-33. Anti-ST2 inhibited rIL-33–induced upregulation of these costimulatory molecules (Fig. 9A). Similar results were observed using splenic DCs from tumor-free mice (Fig. 9B). These data suggest that ST2 mediates costimulatory molecule expression on DCs in response to rIL-33, in line with a requirement for MyD88 for IL-33–induced DC activation. Purified CD11+ DCs from IL-33–stimulated and IL-33+anti-ST2–stimulated DC cultures were compared for their ability to activate Pmel CD8+ T cells when pulsed with gp100 peptides. As expected, IL-33–stimulated DCs exhibited greater T cell proliferation, whereas anti-ST2 inhibited IL-33–induced T cell proliferation (Fig. 9C). To test whether STAT1 activation is through ST2-MyD88–mediated signaling downstream of IL-33, we examined IL-33–mediated STAT1 phosphorylation in DCs from MyD88−/− mice or those treated with anti-ST2. Notably, STAT1 phosphorylation was stimulated by rIL-33 in DCs from WT mice compared with the control. However, this stimulatory effect was abolished by anti-ST2 (Fig. 9D). Moreover, STAT1 phosphorylation was also increased by rIL-33 in WT DCs but not MyD88−/− DCs (Fig. 9D). T cells can express CD11c; therefore, they were depleted from splenocytes using CD90.1 selection prior to purification of CD11c+ cells to exclude their potential role in IL-33–induced DC activation. We further confirmed that IL-33 increased STAT1 phosphorylation only in sorted CD3−CD11c+ WT DCs when they were mixed with MyD88−/− DCs (Fig. 9E), suggesting that IL-33–induced STAT1 phosphorylation is DC intrinsic. Overall, the data indicate that exogenous IL-33 can activate DCs to express costimulatory molecules for maturation through ST2-MyD88–mediated STAT1 signaling.

FIGURE 9.
FIGURE 9.

IL-33, ST2, MyD88, and STAT1 orchestrate activation and maturation of DCs. Purified splenic CD11c+ cells from B16-SIY–bearing mice (A) or tumor-free mice (B) were incubated with the indicated reagents for 48 h, and the frequency of CD40, CD80, CD86, or MHC-II in Gr-1−CD11c+ cells was assessed by flow cytometry. (B) IL-33–induced maturation of naive DCs is also ST2 dependent. (C) rIL-33–treated DCs promote Ag-specific CD8+ T cell proliferation. CD11c+ cells were treated with the indicated reagents for 48 h, as in (B), and then pulsed with gp100 peptides for 2 h. After pulsing, they were cocultured with eFluor 450–labeled naive Pmel CD8+ T cells for 3 d, and proliferation was measured by eFluor 450 dilution. (D) rIL-33–induced STAT1 activation is ST2 and MyD88 dependent. Purified splenic WT or MyD88−/− CD11c+ cells were incubated with the indicated reagents and stained intracellularly for phosphorylated STAT1 by flow cytometry. (E) rIL-33–induced STAT1 phosphorylation is DC intrinsic. Purified CD3−CD11c+ WT and eFluor 450–labeled MyD88−/− DCs were mixed at a 1:1 ratio and treated or not with rIL-33. The next day, cells were analyzed for STAT1 phosphorylation by flow cytometry. WT DCs were gated as e450−, whereas MyD88−/− DCs were CD3−CD11c+e450+. Data (mean ± SEM) are representative of three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001.

Discussion

The roles of IL-33 in tumor growth and tumor immunity have been not well defined. We demonstrate that rIL-33 administration inhibited the growth of established tumors in a CD8+ T cell–dependent manner, consistent with previous studies (21–23). However, our therapeutic approach using rIL-33 is different; Gao et al. (21) used IL-33–transgenic mice, Villarreal et al. (22) used IL-33 DNA vaccination, and Gao et al. (23) used cancer cells expressing ectopic IL-33. More importantly, we are developing the IL-33–based therapy against established tumors rather than targeting very small tumors (0.5–120 mm3 or ones that have only grown for 0–4 d in mice), and we used experimental models of cancer to represent as closely as possible clinical cancers, with the hopes of mimicking a clinical setting.

It is noted that several studies showed an opposite tumor-promoting role for IL-33. One study reported that systemic injection of low levels of rIL-33 increases tumor metastasis (39). Two other studies showed that systemic daily injection of rIL-33 alone (40) or with oncogene (41) for a long duration (>8 wk) promotes tumorigenesis. We speculate that differences in the dosage and duration of IL-33 treatment, as well as the primary target cells of IL-33 in these experimental settings, account for the different in vivo effects. In addition, it was reported that cancer patients tend to have higher IL-33 levels in sera and tumor tissues than do healthy donors (42, 43). Clinically, the levels of circulating soluble ST2 are correlated with tumor burden (44, 45). These data may suggest a role for endogenous IL-33/ST2 signaling in promoting cancer progression (46–48) that is fairly different from the results from our laboratory and other investigators using exogenous IL-33 based on the therapeutic milieu perspective.

One major obstacle of immunotherapy is that the tumor microenvironment renders tumor-infiltrating DCs and DLN DCs dysfunctional in their ability to mount an effective antitumor response. Interestingly, we found that IL-33 therapy increased DC number and upregulated costimulatory molecule (CD40, CD80, CD86, or MHC-II) expression. In ex vivo experiments, DCs from DLNs of IL-33–treated mice were far superior at priming T cells to produce IFN-γ than were those from control groups. Also of note, IL-33 DCs were able to elicit this strong IFN-γ response without the addition of exogenous Ag, indicating that IL-33–treated DCs retain Ag-presenting capabilities, as well as full T cell–priming ability. Most clinically relevant was that we observed this effect in our BRAFV600EPTEN tumors, which express the melanoma-associated Ag gp100 that can serve as an immunotherapy target (49).

We showed that dysfunctional mDCs from tumor-bearing mice can be rescued to cross-prime antitumor CD8+ T cells likely through an ST2–MyD88–STAT1 axis. In support of such a model, it was reported that functional maturation of mDCs requires STAT1 activation and that STAT1 deficiency in DCs leads to impaired costimulatory molecule expression, Ag presentation, and the ability to mount a Th1 responses (50, 51). Moreover, DCs also require STAT-1 phosphorylation for the induction of peptide-specific CTLs (52). Thus, our data suggest that IL-33–induced activation and maturation of mDCs can be regulated, at least in part, by ST2-MyD88–mediated activation of JAK/STAT1 signal-transduction pathways.

Targeting CD40 with an agonist mAb was tested in mice and patients with hematological cancers or solid tumors (53, 54). Several humanized anti-CD40 mAbs have completed phase I clinical trials and are being evaluated in phase II trials. Because rIL-33 therapy resulted in increased expression of costimulation molecules, including CD40, on tumor-associated DCs in vivo, we examined the antitumor effect of agonistic anti-CD40 and rIL-33 combination therapy. The synergistic antitumor activity was observed particularly in the BPS1 model, because anti-CD40 therapy alone failed to inhibit tumor growth. Certainly, the use of other immunotherapeutics, such as checkpoint inhibitors anti–PD-1 and/or anti–CTLA-4, requires further investigation. In contrast, we found that this combination therapy did not trigger severe side effects, including autoimmune disease. Instead, considering the great ability of IL-33 to induce peripheral Treg and MDSC expansion (55), it is likely that combined IL-33 and mAb therapy would increase cancer immunity while reducing autoimmunity. Nevertheless, any sign of autoimmune activity, if present, and the possibility of unintended tissue damage and/or abnormal tissue remodeling resulting from combination therapy also need careful investigation.

Collectively, our results provide new insights into the underlying cellular and molecular mechanisms by which IL-33 regulates antitumor CD8+ T cell immunity to inhibit tumor growth. The immune effects of IL-33 could be highly context dependent and vary in the presence of different cellular targets, phases of immune responses, and model systems. Importantly, we propose how to maximize the therapeutic efficacy of IL-33 administration against established cancer by mechanism-based combinatory approaches.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank the National Institutes of Health Tetramer Facility for providing the Db/gp100 tetramers. We also thank Dr. Xue-Feng Bai for insightful discussions.

Footnotes

  • This work was supported by National Institutes of Health Grant CA149669, the Northwestern Memorial Foundation-Friends of Prentice Grants Initiative, the Specialized Programs of Research Elements Pilot Award (P50 CA090386), the Northwestern University Robert H. Lurie Comprehensive Cancer Center Flow Cytometry Facility, and a Cancer Center Support Grant (NCI CA060553).

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    DC
    dendritic cell
    DLN
    draining lymph node
    4-HT
    4-hydroxytamoxifen
    mDC
    myeloid DC
    MDSC
    myeloid-derived suppressor cell
    MHC-II
    MHC class II
    SIY
    SIYRYYGL
    Treg
    regulatory T cell
    WT
    wild-type.

  • Received June 19, 2015.
  • Accepted November 22, 2016.

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