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Research Articles

Caspase-1 from Human Myeloid-Derived Suppressor Cells Can Promote T Cell–Independent Tumor Proliferation

Qi Zeng, Juan Fu, Michael Korrer, Mikhail Gorbounov, Peter J. Murray, Drew Pardoll, David L. Masica and Young J. Kim
DOI: 10.1158/2326-6066.CIR-17-0543 Published May 2018

Abstract

Immunosuppressive myeloid-derived suppressive cells (MDSCs) are characterized by their phenotypic and functional heterogeneity. To better define their T cell–independent functions within the tumor, sorted monocytic CD14+CD11b+HLA-DRlow/– MDSCs (mMDSC) from squamous cell carcinoma patients showed upregulated caspase-1 activity, which was associated with increased IL1β and IL18 expression. In vitro studies demonstrated that mMDSCs promoted caspase-1–dependent proliferation of multiple squamous carcinoma cell lines in both human and murine systems. In vivo, growth rates of B16, MOC1, and Panc02 were significantly blunted in chimeric mice adoptively transferred with caspase-1 null bone marrow cells under T cell–depleted conditions. Adoptive transfer of wild-type Gr-1+CD11b+ MDSCs from tumor-bearing mice reversed this antitumor response, whereas caspase-1 inhibiting thalidomide-treated MDSCs phenocopied the antitumor response found in caspase-1 null mice. We further hypothesized that MDSC caspase-1 activity could promote tumor-intrinsic MyD88-dependent carcinogenesis. In mice with wild-type caspase-1, MyD88-silenced tumors displayed reduced growth rate, but in chimeric mice with caspase-1 null bone marrow cells, MyD88-silenced tumors did not display differential tumor growth rate. When we queried the TCGA database, we found that caspase-1 expression is correlated with overall survival in squamous cell carcinoma patients. Taken together, our findings demonstrated that caspase-1 in MDSCs is a direct T cell–independent mediator of tumor proliferation. Cancer Immunol Res; 6(5); 566–77. ©2018 AACR.

Introduction

Myeloid derived suppressor cells (MDSCs) are immature myeloid cells that have been functionally defined by their ability to suppress T cells in tumor-bearing mice, as well as cancer patients (1, 2). Depletion of l-arginine and cysteine, increased nitric oxide (NO), upregulation of reactive oxygen species (ROS), peroxynitrates, and multiple cytokines appear to mediate MDSCs' T cell–suppressive function (3–6). As a heterogeneous cell population, however, MDSCs' role as a key mediator of tumor progression has expanded (7–11). These lines of evidence have converged with others who have shown that abnormal myelopoiesis and recruitment of immature myeloid cells intended for inflamed tissue promote cancer cell proliferation by various mechanisms (12–14). We previously showed that STAT3 signaling in human MDSCs can regulate immunosuppressive function of MDSCs, and some of the downstream STAT3-dependent cytokines have been characterized as potentially mitogenic, particularly IL6 (15–17). Other myeloid-derived cytokines, such as IL1β and TGFβ, have been implicated in promoting carcinogenesis (8, 18–20), but these were noted to be primarily T cell–dependent processes. What has not been clear is MDSCs' non–T cell–dependent role in the human tumor microenvironment (TME). Tumors that are refractory to immune checkpoint inhibitors tend to be T-cell poor (21–23), so understanding MDCSs' T cell–independent, procarcinogenic mechanisms are important for the future development of combinatorial immunooncologic trials. Although clinical trials targeting MDSCs and tumor-associated macrophages (TAM) are ongoing (24, 25), how these tumor-infiltrating myeloid cells shape the tumor in a T-cell–poor TME may direct the next wave of combinatorial immunotherapies (26).

The cross-talk between cancer cells and myeloid cells in the context of inflammation-induced tumorigenesis demonstrated that inflammasomes and MyD88 play an important role in murine Apcmin/+ models (27). IL1β and IL18 have a protective role in MyD88-dependent neoplastic development in some of these murine models (28, 29). The NLRP3 inflammasome complex, in some colorectal carcinoma models, can confer an anticarcinogenic effect (30–34). However, other murine models in gastric cancers, B16F10 melanoma, colitis-associated colon cancer, and T-ALL have variously shown opposing protumorigenic activities of inflammasome cytokines (35, 36). Chemotherapy can further confound the picture to activate the NLRP3 inflammasome to promote tumor growth rate (37). In short, the literature points to context-dependent roles of inflammasomes in cancer biology. Many of these inflammation-associated tumorigenesis models have been focused on tumor-intrinsic inflammasomes, and it is unclear how the tumor cell–extrinsic inflammasome/capase-1 axis can regulate tumor cells directly.

To resolve some of this complexity, particularly as to how it relates to human cancer, we focused on the expression and function of caspase-1 in MDSCs from head and neck squamous cell carcinoma (HNSCC) patients. Using both human in vitro and murine in vivo models, we found that caspase-1 from MDSCs can promote T cell–independent, procarcinogenic effects on cancer cells. This report underscores the role of tumor-infiltrating MDSCs as a key regulator of the TME, with both T cell–independent and T cell–dependent procarcinogenic effects, supporting clinical trials targeting caspase-1 and IL1β in solid tumors associated with MDSCs and adds to the body of evidence to potentially warrant changes in the nomenclature of MDSCs.

Materials and Methods

Study approval

Our study was performed in compliance with approved Johns Hopkins Hospital and Vanderbilt University Medical Center Institutional Review Board protocols in accordance with the Belmont Report and US Common Rule. All subjects provided written informed consent prior to participation, and the specimens were deidentified prior to analysis. Included were HNSCC patients undergoing surgery for curative intent. Excluded were those with autoimmune diseases and chronic steroid use. Fresh tumor specimens (typically 1cm3) were harvested from nonmargin areas and processed in PBS within 2 hours of resection.

Reagents and Cells

Cell Proliferation Dye eFluor 450 (eBioscience) was used for cell proliferation assays, according to the manufacturer's protocol. Anti-human CD11b-PE (clone ICRF44), anti-human HLA-DR-FITC (clone G46-6), anti-human CD14–PerCP-Cy5.5 (clone mφP9), anti-mouse Gr-1–Alexa Fluor 700 (clone RB6-8cC5; BioLegend), and anti-mouse CD11b–PE-Cy7(clone M1/70) and Ly-6C-PE (clone AL-21) were obtained from BD Pharmingen. Cell lines used included human head and neck cancer cell lines Cal27, SCC-1, and SCC-25, and human monocytic cell line THP-1. All cell lines were tested free for Mycoplasma. Cell lines were authenticated with GenePrint10 (Promega) through JHH Genetic Resource Core Facility and stored in −70°C in aliquots when not in use. Cells were cultured with RPMI-1640 medium, with 10% FCS, 10% l-glutamine, 10% sodium pyruvate, 10% penicillin/streptomycin, and 0.1% beta-mercaptoethanol. All reagents were from Gibco. When used for cocultures with sorted MDSC, the FCS concentration was 1% to 2%. CD14+ monocytes from healthy non–age-matched volunteers were used for controls. Murine melanoma cell line B16, colon-cancer cell line CT26, and pancreatic cancer cell line Panc02 were obtained from ATCC. MOC1 murine head and neck cancer lines were obtained from Ravi Uppaluri (Dana-Farber Cancer Institute, Boston, MA).

MDSC isolation and flow cytometry analysis

Human MDSCs, characterized as CD11b+CD14+HLA-DRlow/–, were sorted from freshly obtained peripheral blood or tumor tissue from HNSCC patients undergoing surgical treatment at Johns Hopkins Hospital, Johns Hopkins Bayview Medical Center, or Vanderbilt University Medical Center. Peripheral blood mononuclear cells (PBMC) were separated by Ficoll gradient prior to sorting with antibodies against human (h)CD11b, hCD14, and hHLA-DR conjugates, which were stained at 4°C for 30 minutes in dark at 1:50-200 dilution in 50 μL FACS buffer prior to sorting or flow cytometry. Cells were sorted with a MoFlo MLS sorter (Beckman Coulter), as previously described (15). The purity of sorted cells > 90%. All flow data were analyzed under FlowJo software. Human MDSCs were cultured in 2% FCS media.

Murine MDSCs, characterized as Gr-1+CD11b+ or CD11b+Ly6Chigh, were sorted from splenocytes of 1-5 C57BL/6 mice (The Jackson Laboratory) whose tumor (B16, Panc02, MOC1, and MOC2) measured 0.125 cm3. Splenocytes were harvested under sterile condition in RPMI culture media with 10% FCS, filtered with 40-μm cell strainer, and RBCs lysed with ACK buffer (treated for 5 minutes). Fresh splenocytes were stained with antibodies under 4°C in dark for 30 minutes prior to sorting, and these MDSCs were transferred intravenously as indicated in the in vivo tumor models below.

Proliferative index analysis

Sorted MDSCs were cultured in a 37°C incubator with 5% CO2 in 1% to 2% FCS until use, thawed tumor cell lines [human HNSCC cell lines; Cal27, SCC-1, SCC-25 (ATCC, Johns Hopkins Tissue Core) or murine cancer cell lines CT26 or B16] were passaged once, and they were prestained with cell proliferation dye eFlour450 (eBioscience) at a concentration of 5 μmol/L at room temperature for 10 minutes and washed with PBS three times. Tumor cells and sorted MDSCs were cocultured in round-bottom 96-well plates for 3 days. MDSCs/well (1 × 105–5 × 105) were typically used for these assays, with MDSC:tumor cell ratio ranging from 5:1 to 10:1. Stained tumor cells cocultured with peripheral monocytes from healthy individuals were used as controls at the same cell ratios.

The percentage of proliferating tumor cells was measured using the eFlour450 fluorescence signal detected by flow cytometry. Proliferative index was also measured using Ki-67 staining of tumor cells. After coincubation with MDSCs for 3 to 4 days, tumor cells were harvested and fixed with 5 mL 75% ethanol (added dropwise while vortexing) for 3 hours at −20°C, and the fixed tumor cells were stained with FITC-conjugated mouse anti-human Ki-67 (BD Pharmingen) at room temperature for 30 minutes in the dark. Immediately prior to flow cytometry analysis, 10 μL of propidium iodide (PI) staining solution was added to the cells to assess viability. Flow cytometry was used to analyze the stained cells.

Annexin V-FITC (ab14085; Abcam) staining to check for tumor cell apoptosis was assayed according to the manufacturer's instructions. MDSCs' immunosuppression activity was assessed in control MDSCs that were not cocultured with tumor cells at days 3 to 4 to ensure these cells were functionally T-cell immunosuppressive (see Supplementary Fig. S1). Briefly, matched CD4+ T cells from donors were stimulated with CD3/CD28 with or without sorted mMDSCs from HNSCC patients. Either T-cell IFNγ production or CFSE dye dilutional assays were used to confirm the T-cell suppressive activity of mMDSCs.

For cytokine incubation with eFlour450-stained Cal27 cells, we used the following concentrations: rhIL1b (500 pg/mL; R&D Systems), IL18 (500 ng/mL; MBL Medical & Biological Laboratories), IL10 (5 ng/mL; R&D Systems), IL6 (5 ng/mL), IL36 (50 ng/mL; R&D Systems), IL33 (5 ng/mL; R&D Systems), TGFβ (5 ng/mL; R&D Systems), TNFα (100 pg/mL; R&D Systems). Cal-27 cells (1 × 105) were cultured in each well in 96-well, flat-bottom plate. Four to 5 days after the incubation, cells were then collected to run flow cytometry for the proliferation index as described above.

Cytokine array

A Bio-Plex Pro Assays kit (Bio-Rad) was used to measure cytokines on the Bio-Plex system. The supernatant from sorted human MDSCs from peripheral blood or tumor tissue were collected 24 hours after the different treatment conditions. The assay was performed on a 96-well plate. All samples, including standards, were assayed in duplicate or triplicate wells. Standard curve was generated using the Bio-Plex software, and all the values were within the effective detecting limits.

ELISA

Two hundred microliters of the supernatant from each well of cultured human MDSCs under different treatments were collected at different time points for measurement of IL1β and IL18 using ELISA kits without dilutions (eBioscience, BMS618/2, MBL: 7620). Murine MDSCs (5 × 105) were activated with 50 to 100 ng/mL of LPS (Calbiochem) for 24 hours. Each assay had duplicated wells for the standard curve and samples. Concentration was calculated from the standard curve using the recombinant cytokines, per the manufacturer's instructions.

Caspase-1 staining and inhibition

Fluorescent-labeled inhibitors of caspase-1 (FLICA) covalently bind to active caspase enzymes in cells. Caspase-1–specific FLICA 660 Assay kit (ImmunoChemistry Technologies) was used to stain the sorted human MDSCs at 37°C for 1 hour, then washed 2 times with buffer provided in the kit, and counterstained with DAPI, per the assay protocol. FLICA-stained cells (far red) were analyzed with a Hamamatsu Photonics C9100-02 EMCCD camera using the following filter set: Chroma #41137 – Excitation – HQ620/60; Chroma #39187 – Dichroic – Q660LP, and Chroma #40075 – Emission – HQ700/75. The objective used was an Olympus 40× NA 1.4, oil immersion. For caspase-1 inhibition, the caspase-1–specific inhibitor, Z-WEHD-FMK (BD Pharmingen) was added to cultured MDSCs at 80 to 100 μmol/L for 24 hours (for cytokines analysis) or for 5 days (for cell proliferation assays).

Transwell system

HTS Transwell 96-well plates with 1.0-μm polyester membranes were used to determine the effect of MDSCs on tumor cells without cell–cell contact. 105 sorted MDSCs were cultured in the upper chamber, with 105 eFluor450 stained SCC-25 or Cal27 tumor cells in the bottom chamber. Cell proliferative index was determined on days 4 to 6.

In vivo tumor model

Caspase-1–/– mice (The Jackson Laboratory; B6N.129S2 -Casp-1tm1flv or NOD.129S2(B6)-Casp1tm1Sesh Casp4del/LtJ) were housed in Johns Hopkins animal facility under institutional guidelines after animal IACUC protocol was approved. To generate chimeric mice, 5 × 106 bone marrow cells (BMC) from caspase–/– mice or wild-type C57BL/6 mice were adoptively transferred to lethally irradiated (10 Gy) wild-type C57BL/6 mice. Eight weeks after bone marrow transfer, engraftment was confirmed by the presence of CD45+ PBMCs. After engraftment confirmation, B16, MOC1, or Panc-02 tumor cells (1–5 × 105/mouse) were subcutaneously injected into the chimeric mice. Tumors were measured daily or every 2 days with calipers, and mice were sacrificed when the tumors measured 2 cm in the largest diameter. Depleting anti-CD3 (100 μg/mouse from InVivoMab, Lot number: 4773-2/0914 or clone 17A2 from Bio X Cell) were injected intraperitoneally twice a week in some cohorts. Isolated Gr-1+CD11b+ MDSCs (2 × 106–5 × 106; Miltenyi Biotec, Cat#130-094-538) harvested from the spleen of B16- or Panc02-bearing wild-type mice were injected intravenously into caspase-1–/– or wild-type mice with CD3 depletion on days 7, 14, and 21 after tumor injection. In some groups, splenic MDSCs were treated with racemic thalidomide (50 μg/mL; Sigma, Cat#1652500) in RPMI supplemented with 10% FBS at 37°C, 5% CO2 for 2 to 3 hours and then washed with PBS prior to adoptive transfer. Tumor sizes were measured with calipers daily, and tumor volumes were estimated using the formula V (cm3) = 3.14 × [largest diameter × (perpendicular diameter)2]/6.

SiRNA and qPCR

Pre-made inducible lentiviral particles suppressing the MyD88 gene (GenTarget Inc., #LVP 207) were used for siRNA targeting in Cal27 cells. Cal-27 cells at 65% to 70% confluency were transduced with lentiviral construct particles. The RFP reporter gene was used to sort transduced cells by flow cytometry. RFP+ cells were labeled as Cal27-MyD88 KO cells after validation. Total RNA was extracted and reverse transcribed to cDNA with the High-Capacity RNA-to-cDNA Kit (Applied Biosystems). Concentrations of cDNA were measured with Nanodrop 1000 (Thermo Scientific). A Taqman gene expression assay that targets NFkB (Assay ID: Hs00765730_m1) was used to probe for NFκB expression in both the Cal27 and Cal27-MyD88 KO cells. The 18S rRNA was used as the internal control. cDNA (25 ng) was used as the template, and 20 μL reactions were carried out in 96-well plates using the Taqman Universal Master Mix II (2×) as the reaction cocktail. Double delta Ct (ΔΔCt method) analysis on NFkB transcripts was performed to provide its relative quantitation between the two cell lines (38). Assay was carried on the Applied Biosystems Veriti 9902 96-well thermal cycle PCR system. Thermal cycling parameters: 95°C for 10 minutes, denature at 95°C for 15 seconds, and anneal/extend at 60°C for 1 minute, and then repeated for 40 cycles.

TCGA bioinformatics analysis

We compared RNA-Seq by Expectation Maximization (RSEM)-transformed caspase-1 gene expression with progression-free survival (Fig. 5A) and The Cancer Genome Atlas (TCGA)-defined subtype (Fig. 5B). The expression data were from the Broad Firehose (January 28, 2016; https://gdac.broadinstitute.org/), and the survival and subtype data were retrieved from the TCGA Network (39). For each calculation, all samples with available data were used. The Cox proportional hazards ratio, 95% CI (confidence interval), and Wald P value were calculated using the coxph function from the R Survival package (Therneau, “A package for survival analysis in S,” January 1999), where patients were partitioned into two categories using a caspase-1 gene expression Z-score of 1.0 (i.e., patients with a Z-score <1.0 in one category, and patients with a Z-score >1.0 in the second category; Fig. 5A). Significant association between caspase-1 gene expression and the basal versus non-basal TCGA subtypes was calculated as the Wilcoxon rank-sum P value (Fig. 5B).

MDSC gene expression analysis by microarray

Sorted monocytic (CD11b+HLA-DRlowCD14+) and granulocytic (g)(CD11b+HLA-DRlowCD15+) MDSCs from peripheral blood of patients with advanced HNSCC as described (see above) were placed into TRIzol (TRIzol Reagent), and RNA isolation was done using the RNeasy Mini Kit (QIAGEN, www.qiagen.com). RNA quality was confirmed by analysis on the Agilient 2100 Bioanalyzer. Biotin-labeled targets were generated from 20 ng total human RNA using the NuGEN WT-Ovation Pico assay, as per the manufacturer's recommended protocols. The monocytic MDSC microarray, as shown in Fig. 1, was previously described (ref. 40; GEO accession numbers GSE61479 and GSE59047). For the differential gene expression profile between mMDSCs and gMDSCs, Affymetrix Human Gene 1.0 ST microarrays (Affymetrix) were hybridized for 16 hours at 45°C with rotation (60 rpm) as described in Affymetrix's GeneChip Expression Wash, Stain, and Scan User Manual (www.affymetrix.com). Fluorescent signals were determined using Affymetrix's GeneChip Scanner 3000 7G at default parameters described in the manufacturer's GeneChip Expression Analysis Technical Manual. Images were analyzed using the Affymetrix Command Console version 3 (AGCC v3.0), and processed into CEL files at the manufacturer's default settings. RMA normalized log2 signal values were extracted and summarized for gene-level analysis with the Partek Genomics Suite v6.6 (Partek Inc.). In some cases, the gene expression data for gMDSCs and mMDSCs were compared by one-way ANOVA using Partek, and the results were exported for examination and further evaluation to Spotfire DecisionSite (TIBCO Software Inc.). These differential microarray data are archived under GEO accession number GSE112481.

Statistical analysis

The statistical analysis of the results was performed using Prism 6 software or Microsoft Excel v16 software. The ANOVA analysis was used to compare the differences among multiple groups. Two groups were compared using Fisher exact test or Student t test with a two-tailed distribution and calculated at 95% confidence. Differences were considered statistically significant at P < 0.05. Kaplan–Meier curves were compared using the log-rank (Mantel–Cox) test. When present, error bars reflect SEM.

Results

MDSCs with increased inflammasome complex can promote tumor cell proliferation

We sorted monocytic CD11b+CD14+HLA-DRlow/– MDSCs (mMDSC), from head and neck squamous cell cancer (HNSCC) patients and demonstrated MDSC T-cell suppressive activity (Supplementary Fig. S1). Previously, we generated mRNA microarray expression profiles from these sorted MDSCs, and we noted increased expression of inflammasome complex mRNAs that were correlated with increased IL1β and IL18 secretion (Fig. 1A and B; refs. 15, 40). To characterize the functional significance of the inflammasome signaling in human mMDSCs, we modeled the human TME using cocultures of multiple HNSCC tumor cell lines (Cal27, SCC-1, SCC-25) with sorted mMDSCs to examine whether MDSCs can directly regulate the proliferation of tumor cells. When fluorescently labeled cancer cells were cocultured with mMDSCs without T cells, the proliferation index of the tumor cells was significantly enhanced in all HNSCC lines assayed without any changes in apoptosis (Fig. 1C, left; Supplementary Fig. S2). Most of the MDSCs sorted from the 21 HNSCC patients showed this ability to promote tumor cell proliferation, whereas monocytes from healthy donors did not alter tumor cell proliferation (Fig. 1C, top right). The protumorigenic MDSC response was dependent on the ratio of MDSCs to tumor cells (Fig. 1C, bottom right). MDSCs also increased the tumor cells' expression of Ki-67, a marker of cell proliferation, corroborating the dye dilutional assays (Fig. 1D).

Figure 1.
Figure 1.

Inflammasome complex expression is increased in human MDSCs and can directly increase proliferation of tumor cells in vitro. A, Relative mRNA expression profile of inflammasome complex genes via an MDSC microarray. CD3 transcript expression was used as the relative negative control to generate folds changes (listed on the y-axis). B, Secretion of IL1β and IL18 from sorted human MDSCs using Bio-Plex. Control: culture media only. C, Sorted MDSCs from HNSCC patients and CD14+CD11b+ monocytes from healthy donors (controls) were cocultured with HNSCC cancer cell lines (T)—SSC-1, SCC-25, Cal27—prelabeled with cell proliferation dye eFluor 450. Dye-dilutional shifts of live tumor cells were used to quantitate fold change of proliferating cells. Top left, Representative plot showing proliferation of tumor cells with MDSCs or control monocytes. Bottom left, Summarizes proliferation changes for 3 HNSCC cell lines. Top right: Proliferation changes of sorted mMDSCs (CD14+CD11b+HLA-DRlo) from advanced HNSCC patients (N = 21, error bars are the SEM of fold changes from each biospecimens) or control monocytes cocultured with Cal27 cells. Bottom right: Different ratios of cancer cells to MDSCs were used to show dose–response effect on tumor proliferation. Cal27:MDSC = 1:5 or 1:10. T = Cal27 only. D, Expression of Ki-67 was analyzed by flow cytometry in tumor cells after incubation with MDSCs or control monocytes. Histograms represent Ki-67 expression in different groups. T: MDSC = 1:5. Control groups, Isotype control antibody and control monocytes from healthy individuals. **, P < 0.01; ***, P < 0.001. Each of the in vitro experiments was repeated at least 3 times independently. Error bars in B represent SEM from triplicates.

To investigate whether the upregulated inflammasomes in MDSCs were related to their promotion of tumor cell proliferation, we treated human mMDSCs with the caspase-1–specific inhibitor, Z-WEHD-FMK, which reduced the secretion of IL1β and IL18 prior to the coincubation assay (Fig. 2A). A caspase-1–specific FLICA assay was used independently to corroborate that caspase-1 was active in situ in the sorted human MDSCs (Fig. 2B). When the caspase-1 inhibitor, Z-WEHD-FMK, was added to the MDSCs prior to coculture, we found that MDSCs decreased their ability to induce tumor cell proliferation (Fig. 2C).

Figure 2.
Figure 2.

Inhibition of caspase-1 activity reduces MDSCs' ability to promote carcinogenesis. A, IL1β and IL18 production (ELISA) by sorted HNSCC patient mMDSCs (CD14+CD11b+HLA-DRlo) with and without inhibition of caspase-1 using Z-WEHD-FMK. **, P < 0.01; *, P < 0.05. Control, Monocytes from healthy individuals without inhibitor. Bar graphs are representative of at least 3 repeats of experiments with duplicate wells. B, FLICA (fluorescent labeled inhibitor of caspase-1), covalently binds to active caspase-1, was used to visualize caspase-1 activity in mMDSCs. Nuclei were counterstained with DAPI (black arrow). Caspase-1 staining is far-red FLICA fluorescence (white arrow). Pictures shown in 40× and 100× magnifications. Scale bar, 100 μm at 40×, and 250 μm in 100×. C, Proliferation (%) of Cal27 with and without Z-WEHD-FMK. Control group: Cal27 cultured with monocytes from healthy individuals. *, P < 0.05. Each of the in vitro experiments was repeated at least 3 times independently. Error bars in A and C represent SEM from triplicates.

Paracrine mitogenic activity is higher in monocytic MDSCs

When we compared the mitogenic induction between sorted gMDSCs (CD15+CD14–HLA-DRlow/–) to the mMDSCs from the same patients, we noted that mMDSCs had greater ability to promote tumor proliferation (Fig. 3A). When we compared the relative expression profiles between mMDSCs and gMDSCs in a new microarray profile, mMDSCs had greater expression of inflammasome complex mRNAs than gMDSCs (Fig. 3B). We reasoned that if the promotion of tumor proliferation is dependent on caspase-1/IL1β/IL18 signaling pathway, this phenotype would be a paracrine-dependent mechanism, and cell–cell contact would not be necessary. We performed the tumor proliferation assay in a Transwell assay, and we noted that the presence of mMDSCs in the test chamber increased the proliferation index of tumor cells in the other chamber (Fig. 3C, P < 0.001). We also showed that supernatants from human mMDSCs phenocopied the enhanced proliferative index of HNSCC cells coincubated with MDSCs (Fig. 3D, right). When we followed the proliferative index of Cal27 with the incubation of recombinant human cytokines expressed by MDSCs, enhanced proliferation was noted when IL1β, IL18, IL6, and IL10 were combined, whereas IL1β and IL18 individually only had about a 1.3-fold effect (Supplementary Fig. S3).

Figure 3.
Figure 3.

MDSCs' promotion of tumor proliferation is differentially enhanced in monocytic MDSCs and granulocytic MDSCs and is not dependent on cell–cell contact. A, Matched monocytic MDSCs were compared with granulocytic MDSCs (CD15+CD14–HLA-DR–) in their ability to promote tumor proliferation. Assays were replicated 5 times, and a representative result is displayed. *, P < 0.05. B, Volcano plot of the differential microarray analysis of inflammasome complex genes between mMDSCs and gMDSCs. C, Transwell assay to assess whether cell–cell contact was required for proliferation promotion. Transwell upper chamber: Sorted human MDSCs: Transwell bottom chamber: Seeded eFluor405-stained Cal27 cells. Fresh RPMI plus 2% serum without MDSCs were used as controls (in the upper chamber). Proliferative index was determined by dye dilutional assay using flow cytometry. Representative plot also shown (top). ***, P < 0.001. D, Supernatant from mMDSCs were added to eFluor405-stained SCC-1 cells, and the proliferative index was determined as noted above. Fresh RPMI (with 2% serum) with monocytes from healthy individuals was used as controls. Dot plots (top) represent the dye dilutional analysis, and the bar graph (bottom) shows the fold changes of tumor proliferation. **, P < 0.01. Each of the in vitro experiments was repeated at least 3 times independently. Error bars in A, C, and D represent SEM from triplicates.

Myeloid caspase-1 directly promotes carcinogenesis

Because the in vitro cell mixing experiments did not replicate the complexity of the TME, we directed our attention toward in vivo characterization of myeloid caspase-1, in the context of a syngeneic tumor model. In order to probe whether MDSC caspase-1 can directly promote tumor proliferation in vivo, we used chimeric C57BL/6 mice adoptively transferred with BMCs from caspase-1 null or wild-type control mice. We initially showed that murine Gr1+CD11b+ MDSCs expressed IL1β and IL18 (Supplementary Fig. S4), and its coculture with B16 or CT26 tumor cells promoted caspase-1–dependent tumor cell proliferation as we observed with human mMDSCs (Fig. 4A). In chimeric mice with caspase-1 null hematopoietic cells, B16 tumor growth was significantly blunted, regardless of whether CD3+ T cells were depleted or not (Fig. 4B, top left). The T cell–independent effect of caspase-1 signaling was more prominent in this subcutaneous tumor injection model. We repeated the experiment with Panc02 pancreatic cancer cells (Fig 4B, top right), as well as MOC1 oral cavity tumor cells (Supplementary Fig. S5) with the same results. When wild-type Gr1+CD11b+ MDSCs harvested from B16-bearing mice were adoptively transferred into caspase-1 null chimeric mice after tumor injection, the protective effect of absent caspase-1 signaling was reversed (Fig. 4B, bottom left). To test whether pharmacologic blockade of MDSC inflammasome signaling can specifically phenocopy the caspase-1 null chimeric mice, we treated wild-type MDSCs with caspase-1 inhibiting thalidomide prior to their adoptive transfer into tumor-bearing mice that had caspase-1 deficiency and were T-cell depleted (41). Although the wild-type MDSCs restored their protumorigenic effect, thalidomide-treated MDSCs failed to increase the protumorigeneic tumor growth rate of B16 in chimeric mice (Fig. 4B, bottom right). Together, these results show that caspase-1 activity in MDSCs could directly promote in vivo tumor growth, independent of T cells.

Figure 4.
Figure 4.

MDSC caspase-1 directly promotes carcinogenesis. A, Murine Gr1+CD11b+ MDSCs (produced IL1β and IL18, Supplementary Fig. S3) were cocultured with CT26 and B16 cells (prestained with eFluor450), and the fluorescence dilution of tumor cells were analyzed. Each experiment was performed in triplicate and repeated 3 times. B, Tumor growth experiments in vivo were performed with chimeric C57BL/6 mice adoptively transferred with BMCs from caspase-1–/– mice (KO) or wild-type (WT) mice. BMC reconstitution was confirmed at 8 weeks. 1 × 105–3 × 105 B16 (top left) or Panc02 (top right) cells were subcutaneously injected to the right flank area and the injected tumor was measured daily. Bottom graphs: Tumor volume. CD3+ T cells were depleted via antibody depletion i.p. twice a week in chimeric C57BL/6 mice adoptively transferred with BMCs from caspase-1 KO or WT mice. Wild-type Gr-1+CD11b+ MDSCs (enriched from splenocytes of tumor-bearing wild-type mice) were transferred into tumor-bearing chimeric mice (bottom left). MDSCs were also treated thalidomide (50 μg/mL) prior to MDSC transfer in some groups (bottom right). *, P < 0.05; **, P < 0.01.

Caspase-1 expression in the TME correlates with worse prognosis

To investigate the role of caspase-1 as a potential prognostic marker in HNSCC patients, we correlated caspase-1 gene expression with survival and subtype using data from a TCGA HNSCC patient cohort (Fig. 5; ref. 39). Elevated caspase-1 expression was significantly associated with decreasing progression-free survival in TCGA HNSCC patients (Fig 5A). Similarly, when we probed for caspase-1 expression in the basal subtype, which has previously been associated with worse clinical outcomes with relative absence of T-cell infiltration 42, 43), caspase-1 expression was significantly higher in the basal subtype than in the nonbasal subtypes (Fig. 5B).

Figure 5.
Figure 5.

Increased caspase-1 expression is associated with worse prognosis in a TCGA HNSCC dataset. Analysis of TCGA data for caspase-1 gene expression in HNSCC patients. A, Kaplan–Meier showing progression-free survival. Of 46 patients for whom CASP1 gene expression and progression-free survival data were available, nine were defined as overexpressing CASP1 (Z-score ≥ 1.0; red line) and 36 were defined as have wild-type expression (Z-score < 1.0; blue line). B, Box plot of caspase-1 expression in basal and nonbasal subtypes. Boxplots: Interquartile range (IQR; box height) with the upper horizontal line showing the 75% quantile, lower showing the 25% quantile, and center line representing the median. The vertical line (whisker) extends below the lower quantile (LQ) to minimum of LQ – (1.5 * IQR) and extends above the upper quantile (UQ) to a maximum of UQ – (1.5 * IQR). Data points show patients outside of the IQR. A Z-score cutoff of ≥1.0 was used to define patients as overexpressing caspase-1 (see Materials and Methods). CI: confidence interval.

MDSCs' promotion of tumor proliferation is dependent on tumor cell MyD88 signaling

Because both IL1β and IL18 signaling results in activation of the tumor MyD88 signaling pathway to induce oncogenic transcription factors NFkB and AP-1 (27, 44), we inquired whether MDSCs' stimulation of MyD88 signaling is partly responsible for the enhanced mitogenic signaling in HNSCC cells. We suppressed the MyD88 gene expression in Cal27 HNSCC cell line with siRNA that also expressed an RFP reporter gene, and we sorted the RFP-expressing Cal27 cells. MyD88 knockdown in Cal27 cells showed significantly lower NFkB transcripts, as indicated by the higher number of NFkB qPCR cycling (Fig. 6A). When we compared the proliferation index of wild-type Cal27 cells and the MyD88 suppressed Cal27 cells cocultured with MDSCs, the MDSC-dependent promotion of tumor cell proliferation was significantly blunted in the MyD88-suppressed tumor cells in vitro (Fig. 6B). We reasoned, therefore, that tumor cells deficient in MyD88 signaling would not be effected by myeloid caspase-1 signaling in vivo. Thus, we compared the tumor growth rate of B16 cells and B16 MyD88 knockdown cells in wild-type mice and found that MyD88 suppression significantly decreased the growth rate of B16 in vivo (Fig. 6C). However, using chimeric mice adoptively transferred with caspase-1 null BMCs under T cell–depleting conditions, MyD88 signaling in the B16 tumor cells resulted in no differential tumor growth rate in vivo compared with control tumor cells with transduction of scrambled shRNA (Fig 6C). The blunted tumor growth rates attributable to myeloid caspase-1 signaling seen previously in Fig. 4 were not found in these tumors cells lacking MyD88/NFkB oncogenic signals.

Figure 6.
Figure 6.

Caspase-1 from myeloid cells induces tumor proliferation via MyD88 oncogenic signaling. A, Cal27 cells were transduced with MyD88 siRNA lentivirus (MyD88 knockdown, KD) and probed for NFkB expression via qPCR. Left, Transduction efficiency of the reporter RFP gene in Cal27 with and without MyD88 KD. Right, Relative NFkB gene expression between the control Cal27 cells (transduced with scrambled lentivirus) and MyD88 knockdown Cal27 cells (T-MyD88 KD). Higher ΔΔCt value represents greater cycling which inversely reflects low expression of gene. B, Sorted human MDSCs were cocultured with Cal27 cells with and without MyD88 knockdown, and tumor cell proliferation was assessed. Representative histogram plots of Cal27 proliferation under different conditions shown (left) in addition to fold change in proliferation (right). **, P < 0.01. C, In vivo tumor growth curve of B16 and MyD88 KD-B16 cells in caspase-1 null chimeric mice (all with T-cell depletion). *, P < 0.05; n.s., not significant. Each of the in vitro experiments was repeated at least 3 times independently. Error bars in B represent SEM from triplicates.

Discussion

Depending on the preclinical models used, the role of the inflammasome in carcinogenesis has been contradictory and context dependent (27, 35). In one colorectal metastasis model, the inflammasome complex was found to suppress metastasis via NK cell activity (45), and others showed that NLRP3 signaling in hematopoietic compartment can be protective against tumorigenesis through bone marrow chimera studies (31, 33). In other models, however, NLRP3 null mice had reduced lung metastasis with B16-F10 tumor cells (46). These examples highlight the complex association between the inflammasome complex and cancer, which were have been reviewed in detail (47–49). How these various context-dependent mechanisms of the inflammasome complex in murine models are pertinent to human cancer biology is still unclear. Hence, clinical development of inflammasome targeting agents has been somewhat hindered by these conflicting preclinical murine results. However, an intriguing correlative finding was noted in atherosclerotic patients treated with canakinumab (blocks IL1β; refs. 50, 51). Although canakinumab effectively reduced myeloid dependent atherosclerotic disease, the treated group also showed surprising reduction in lung cancer incidence and cancer-associated mortality. Combined with our findings, translational development of IL1β or inflammasome blocking agents in human malignancies is warranted.

Although the TME is infiltrated with both lymphocytic and myeloid cells, there are a subset of aggressive carcinomas that are T-cell poor and may be amenable for MDSC targeting (21). Interestingly, these “noninflamed” tumors have been reported to be less responsive to checkpoint inhibitors, and the ability to inhibit protumorigenic myeloid cells will be an important clinical strategy going forward in immune oncology (52). Our previous report in preclinical MOC head and neck tumor model with higher MDSC tumor infiltration relative to tumor-infiltrating lymphocytes was more responsive to immune checkpoint inhibition with the addition of MDSC-depleting antibody (53). Rilonacept and canakinumab are two examples of IL1β targeting agents that are safe in early clinical trials for chronic inflammatory diseases, so clear clinical pathways to translate inflammasome targeting agents into immune-oncological trials exist (49, 50, 53). Our demonstration that MDSC caspase-1 directly promoted NFkB-dependent tumor proliferation also offers potential biomarker correlative analysis on the tumor intrinsic MyD88/NFkB signaling that may be associated with clinical response with these anti-inflammasome agents.

With regard to clinical development, several clinical trials that target myeloid chemokine receptors (www.clinicaltrials.gov) are under way. Although our findings cannot answer how to best target MDSCs, pharmacologic use of inflammasome blocking agents, as shown by our experiments with thalidomide, are supported by our data. The potential problem with chemokine receptor blocking strategies is that different types of MDSCs and other immunosuppressive myeloid cells may have differential recruitment pathways. We previously showed that CXCR2 may be responsible for granulocytic MDSC recruitment, whereas CSF1R primarily recruits monocytic MDSC (53). Others have shown that IL1β from epithelial cell can recruit MDSCs into the tumor (46). We, therefore, support a global pharmacologic targeting of the inflammasome pathway in tumor cells, as well as myeloid cells, to potentially induce greater antitumor responses.

For the downstream mechanism of action of caspase-1 in the TME, the literature points to discordant effects of IL1β and IL18, the two primary downstream cytokines from caspase-1 activation. In our limited cytokine-Cal27 incubation experiments, a significant tumorigenic response to these cytokines, primarily when the cytokines were combined, was seen. In general, IL1β has been linked to a tumor induction and progression, whereas IL18 has been associated with NK cell–dependent protection against tumors (44, 54–56). From our adoptive transfer experiments with caspase-1 null mice, we hypothesize that other caspase-1 dependent secretome factors or IL1β may prevail over the potential protective impact of IL18 induction of NK cells in the TME. There are potentially other mechanisms that may explain our in vivo data, such as caspase-1's promotion of tumor proliferation may be one of many T cell–independent mechanism of carcinogenesis. The dramatic antitumor effect in our chimeric mice suggests that other downstream effectors of inflammasome complex, such as pyroptosis, may also regulate the TME. Our findings offer an avenue of research assessing downstream effectors of myeloid inflammasomes that can shape both the primary tumor and the premetastatic niche in both the draining lymph nodes and distant organ sites (26).

Although we focused on caspase-1 in MDSCs, their activity on macrophages and other myeloid cells that infiltrate the tumor will require further investigation. Given the plasticity of MDSCs in the TME and their tendency to differentiate ex vivo, the ability to differentiate tumor-associated MDSCs from other myeloid cells may be difficult. Although early reports show that immature VEGF+ myeloid cells may help establish the premetastatic niche and the discoveries that STAT3 in myeloid cells can promote metastasis (57, 58), tumor-infiltrating myeloid cells, such as MDSCs, may become an important regulator of metastasis. These previously reported links between MDSCs and metastasis may explain how myeloid caspase-1 signaling is associated with overall survival in the TCGA database. Although most of these works have focused on how these immunosuppressive myeloid cells regulate the intermediary immune cells to render cancer more metastatic, we presented evidence in the human system that MDSCs could have a direct mitogenic response, which may be relevant in the context of the premetastatic niche. Such a direct procarcinogenic mechanism has been advanced in murine models of lung metastasis, which shows MDSCs may promote epithelial–mesenchymal transition (26). Although our in vitro system did not account for T cells, NK cells, and stromal cells in the TME, we argue that the immunosuppressive mechanism of MDSCs in the TME must also account for the basal oncogenic signals that MDSCs can provide for the tumor cells. Lastly, our report further supports the need to redefine the functional definition of MDSCs (59).

Disclosure of Potential Conflicts of Interest

No potential conflicts of interest were disclosed.

Authors' Contributions

Conception and design: Q. Zeng, J. Fu, P.J. Murray, Y.J. Kim

Development of methodology: Q. Zeng, J. Fu, M. Gorbounov, Y.J. Kim

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): Q. Zeng, J. Fu, M. Gorbounov, Y.J. Kim

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): Q. Zeng, J. Fu, M. Korrer, P.J. Murray, D.L. Masica, Y.J. Kim

Writing, review, and/or revision of the manuscript: Q. Zeng, J. Fu, M. Korrer, M. Gorbounov, P.J. Murray, D. Pardoll, Y.J. Kim

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): J. Fu, Y.J. Kim

Study supervision: Y.J. Kim

Acknowledgments

The authors thank the NIH for support of this research (NIH R01CA178613). The authors also thank the Bloomberg-Kimmel Institute for Cancer Immunotherapy and the Baker family for support.

We would like to acknowledge Dr. Ravi Uppaluri (Dana-Farber Cancer Center) for providing the MOC1 HNSCC cell lines. We would also like to thank Conover (Connie) Talbot Jr (Johns Hopkins School of Medicine) for assisting the bioinformatics of the microarray analysis.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Footnotes

  • Note: Supplementary data for this article are available at Cancer Immunology Research Online (http://cancerimmunolres.aacrjournals.org/).

  • Received September 25, 2017.
  • Revision received January 2, 2018.
  • Accepted March 26, 2018.
  • Published first April 13, 2018.

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