The Expression of Adenosine A2B Receptor on Antigen-Presenting Cells Suppresses CD8⁺ T-cell Responses and Promotes Tumor Growth

Adenosine is present in the extracellular space and functions as a signaling molecule by engaging four different adenosine receptor subtypes; A1, A2AR, A2BR, and A3 (1). Among these, A2AR and A2BR are implicated in the signaling that resolves inflammation and promotes tissue repair (2). Adenosine-mediated tissue-healing responses can help tumors escape immune recognition and disseminate to other tissues (3–5). Therefore, identifying the roles of adenosine receptor subtypes in different subsets of immune cells is needed for developing rational strategies targeting adenosine signaling as a potential cancer therapy.

A2BRs are expressed in both immune and nonimmune cells and have pleotropic protumoral effects. A2BR expression in tumor cells is implicated in decreased adherence, increased survival, and increased metastasis in both immune-deficient and proficient settings (4, 6–8). A2BR is also expressed by immune cells (9, 10), and A2BR expression is particularly increased in antigen-presenting cells (APC) upon activation (11). Targeting A2BR by genetic deletion or by pharmacologic blockade slowed the growth of several syngeneic tumors by activation of T cells (4). In addition, promotion of tumor growth by A2BR signaling associates with expansion or activation of suppressive myeloid cells (12–14) and transfer of myeloid suppressor cells reverses the enhanced adaptive immune responses during A2BR blockade (12). However, it is not known if cell-intrinsic A2BR signaling by T cells versus APCs plays a major role in increasing tumor growth. Also, the impact of activating A2BRs on myeloid cells and APCs on lung colonization of tumors remains elusive.

Here, we confirmed that A2BR deletion delayed growth of a number of syngeneic ectopic solid tumors in a T-cell–dependent manner. Experiments utilizing adoptive transfer of wild-type (WT) versus A2BR^−/− T cells to tumor bearing hosts or growth of tumors in mice reconstituted with a mixture of WT and A2BR^−/− bone marrow (BM) cells demonstrated that T-cell A2BRs had a very limited role in A2BR-mediated suppression of tumor-associated T cells. Adoptively transferred tumor antigen-specific CD8⁺ T cells were more cross-primed in tumor bearing A2BR knockout (KO) hosts than that of WT hosts. A2BR deletion intrinsically favored accumulation and activation of myeloid APCs and Gr1⁺ MDSCs in the tumor microenvironment. Both CD11c and myeloid deletion of A2BR delayed growth. However, lung colonization of tumors was only inhibited by myeloid deletion of A2BR, suggesting monocytes and other Gr1⁺ cells were important for A2BR-mediated promotion of lung dissemination. Finally, acute pharmacologic blockade of A2BR before adoptive transfer of tumor antigen–specific T cells improved antitumor efficacy compared with control or the single treatments. Thus, these findings suggested clinical potential for A2BR blockade to improve current cancer immunotherapies.

Animal experiments were approved by the Animal Care and Use Committee of the La Jolla Institute for Allergy & Immunology (La Jolla, CA) and the institutional animal use committees of Northwestern University (Chicago, IL) and Bilkent University (Ankara, Turkey). B16F10 cells stably expressing luciferase (B16-Luc) were obtained from Caliper Life Sciences and cultured in R5F (RPMI1640 medium containing 10% heat-inactivated FBS, 2 mmol/L l-glutamine, 1 mmol/L sodium pyruvate, 50 U/mL penicillin, 50 mg/mL streptomycin). Cell lines from Caliper Life Sciences were tested for being pathogen free. Dr. D. Theodorescu at University of Colorado (Aurora, CO) kindly provided us with the mouse urethelial carcinoma cell line MB49 originally produced by Dr. T. Ratliff of Purdue University (West Lafayette, IN). MB49 cells were confirmed to be mouse origin and tested negative for evidence of cross-species contamination and pathogen contamination by IDEXX BioResearch. All cell lines were passaged less than 10 times after initial revival from frozen stocks. Murine Lewis lung carcinoma (LLC1) cells were purchased from ATCC (CRL-1642). LLC1 cells expressing OVA (LLC1-OVA) were generated as described previously (15). For all experiments, tumor cells were recovered from frozen aliquots and cultured for 1 to 2 weeks prior to inoculation of mice. All the cell lines were routinely tested for Mycoplasma infections by culture and DNA stain and maintained in complete medium composed of RPMI1640 with 5% FBS, but have not been reauthenticated in the past year. Six-week-old C57BL/6 Rag1^−/−, CD45.1 and CD90.1, LysMCre mice [B6.129P2-Lyz2tm1(cre)Ifo/J), CD11c-Cre (B6.Cg-Tg(Itgax-cre)1-1Reiz/J] and Rag1^−/− mice were purchased from Jackson Laboratories. A2BR^−/− mice were provided by Dr. Michael R. Blackburn (The University of Texas Health Science Center, Houston, TX) or by Katya Ravid of Boston University (Boston, MA). Dr. Hans Schreiber (University of Chicago, Chicago, IL) provided the 2C transgenic mice, SIYRYYGL peptides and MC38, B16-SIY cell lines. Adora2b^f/f mice were generated as described previously (16) and crossed with LysMCre^−/+ mice. The 10- to 12-week-old mice with adenosine receptor deletions used in this study were congenic to C57BL/6 and were created as described previously: A2AR^−/− (17), A2BR^−/− (18, 19). Antibodies used for flow cytometry are listed in Supplementary Table S1. Cytokines were used for the study is listed as Supplementary Table S2. All adenosine receptor agonists and antagonists were purchased from Tocris Bioscience (Supplementary Table S3).

B16-SIY, B16-Luc, LLC1, LLC1-OVA, or MC38 cells (1 × 10⁶) and MB49 cells (1 × 10⁵) in 100 μL of PBS were injected subcutaneously. For A2BR blockade in vivo, 10 days after tumor cell injection, mice were injected intraperitoneally by MRS1754 in 100 μL of 0.1% DMSO (2 mg/kg) once daily. The tumor volume was determined by calipers at 2- to 3-day intervals. Tumor volumes were measured along 3 orthogonal axes (a, b, and c) and calculated as (a*b*c)/2.

BM chimeric mice were generated as described previously (20). Briefly, A2BR^−/− KO mice and WT mice were exposed to 10 Gy total body irradiation using a Cesium-137 Mark I irradiator (J.L. Shepherd Inc, San Fernando, CA). BM cells from the femur and tibia of matched A2BR^−/− mice and WT mice were harvested under sterile conditions. Irradiated recipient mice received 10⁷ BM cells in 100 μL of PBS intravenously. Mice were housed for 8 weeks after BM transplantation before experimentation. BM reconstitution of Rag1^−/− mice with BMs from WT and A2BR^−/− mice was performed as described previously (21). Briefly, mice 6–12 weeks of age were fasted for 24 hours and then lethally irradiated (2 × 450 Rads for Rag1^−/− and 2 × 500 Rads for C57BL/6 recipients) using RS2000 Biological irradiator (Rad Source Technologies Inc.). After the second radiation exposure, a 1:1 mixture of 5–10 × 10⁶ BM cells from donor mice (WT and A2BR^−/−) in 100 μL of PBS were injected intravenously through the retro-orbital venous sinus. Mice were treated with antibiotics starting 3 days before tumor implantation up until 2 weeks after radiation. We injected 10⁵ MB49 bladder carcinoma cells in 100 μL of PBS subcutaneously into the right flanks 10 weeks after reconstitution of the BM. Single-cell suspensions from spleens and tumors were analyzed by cytofluorometry to determine the proportion of T cells, NK cells, and myeloid cells 7, 14 and 21 days after tumor inoculation as described below.

Tumor tissues were minced into small pieces and digested in Collagenase IV (1 mg/mL; Roche) and DNase I (20 μg/mL; Roche) at 37°C for 30 minutes, and filtered by sequential pressing through 100- and 40-mm cell strainers. After red blood cell (RBC) lysis using RBC lysis buffer (BioLegend, catalog no. 420301) according to the manufacturer's instructions, cells were washed and resuspended in R10F and counted in a Z2-Coulter particle counter (Beckman Coulter). For BM-derived macrophages (BM-DM), cells were detached with Accutase solution (Innovative Cell Technologies, catalog no. AT104) 6 hours after 100 ng/mL LPS (Invivogen, catalog no. tlrl-smlps) treatment in the presence or absence of 1 μmol/L NECA (Tocris Bioscience, catalog no. 1691). Single-cell suspensions were preincubated for 10 minutes in 100 mL FACS buffer with antibody to block Fc receptors. Each sample tube received 100 mL fluorescently labeled antibody mixture (list of antibodies are indicated in Supplementary Table S1) and was incubated for 30 minutes at 4°C in the dark. For intracellular cytokine staining, tumor samples were restimulated with 10 ng/mL PMA, 100 ng/mL ionomycin (Sigma-Aldrich), and Golgi Plug (eBioscience) for 5 hours at 37°C. Cells were fixed and permeabilized after surface staining and incubated for 25 minutes at 4°C in 100 mL permeabilization/washing buffer (BioLegend, catalog no. 421002) containing 1:100 anti–IFNγ for tumor samples or anti-TNFα and anti-IL10 for BM-DM samples. After a subsequent wash, cells were resuspended in 350 mL FACS buffer. Cells were analyzed using an LSR II equipped with four lasers and FACS Diva software (BD Biosciences). Live/dead fixable yellow (Invitrogen, catalog no. L34959) was used to exclude dead cells according to the manufacturer's instructions before analysis. Flow cytometry data were analyzed using FlowJo software (9.0.1 version; Tree Star) or Novocyte software (1.4.1. version; Acea Biosciences). Gating strategy for all the tested subpopulations and activation markers are listed in Supplementary Fig. S1.

For metastasis analysis, 3 × 10⁵ B16-Luc in 100 μL PBS were injected intravenously into the tail vain and luciferase activity was measured one and two weeks after the injection of cancer cells by injection of 1 mg d-Luciferin (Caliper Life Sciences) in 100 μL PBS followed by IVIS Imaging (Caliper Life Sciences). Images were taken within 10 minutes of luciferase injection. Data were collected for 1 minute and represented as photons/second. After measuring luciferase activity, lungs were removed, and pictures from representative lungs were taken to observe metastatic dark spots. Lungs were also weighed to validate changes in overall metastatic tumor burden per group in LysMCre^−/+A2BR^f/f versus littermate controls.

IL12 ELISA was performed after exposing LPS-stimulated BM-derived dendritic cells (BM-DC) to various adenosine receptor antagonists (listed below and in Supplementary Table S3) in the presence of NECA or to various receptor subtype–specific adenosine receptor agonists. For cytokine detection IL12p40 ELISA kit (BioLegend, catalog no. 431601) was used by following manufacturer's instructions. For ELISA experiment, 100 nmol/L of following receptor subtype–specific agonists and antagonists from Tocris Biosciences were used: A1R agonist, 2′-MeCCPA; A2AR agonist, CGS 21680 hydrochloride; A2BR partial agonist, BAY 60-6583; A3R agonist, 2-Cl-IB-MECA; A1R Antagonist, PSB36; A2AR Antagonist, SCH 58261; A2B Antagonist, PSB 603.

Splenic CD8⁺ T cells from CD45.2⁺ 2C WT CD90.1⁺ or 2C A2BR^−/− CD90.1⁻ mice were selected with a CD8⁺ T Cell Enrichment Kit (StemCell Technologies, catalog no. 19853). For CFSE labeling, cells at 10⁶ per mL were stained with CellTrace CFSE staining solution (Thermo Fisher Scientific, catalog no. C34554) for 20 minutes in a 37°C water bath according to the manufacturer's instructions. A total of 2 × 10⁶ purified CFSE-labeled T cells in 100 μL of PBS were injected intravenously into CD45.1⁺ hosts through the retro-orbital venous sinus 1 day prior to B16 or B16-SIY injection. Three days later, the proliferation (CFSE dilution), intracellular IFNγ⁺ production and activation (measured by expression of CD69 and CD44) of transferred T cells (gated on CD8⁺CD45.1⁻ CD45.2⁺) in spleens were determined by flow cytometry. For adoptive T-cell therapy, splenic CD8⁺ T cells were selected from 2C mice with a CD8⁺ T Cell Enrichment Kit (StemCell Technologies, catalog no. 19853) and stimulated with 0.5 μg/mL SIY peptides, 1 μg/mL anti-CD28, and 10 ng/mL IL2 in CR10, that is, RPMI1640 supplemented with 10% FCS (Life Technologies, catalog no. 26140079), 100 IU/mL penicillin, 100 μg/mL streptomycin, 2 mmol/L l-glutamine, 25 mmol/L HEPES buffer, and nonessential amino acids for 3 days. Prestimulated T cells were injected intravenously at 5 × 10⁶ in 100 μL of PBS per mouse through the retro-orbital venous sinus into B16-SIY tumor–bearing mice on day 1 or day 13. To examine the role of A2BR on regulatory T cells (Treg), tumor-associated or splenic CD4⁺CD25^hi Tregs from WT or A2BR^−/− were sorted by BD Aria. For the colitis model as described previously (22), mouse splenocytes were stained with anti-CD45RB, anti-CD25, and anti-CD4, and sorted by BD Aria for CD4⁺CD45RB^hi and CD4⁺CD25^hi T cells (considered as Tregs), respectively. A total of 4 × 10⁵ sorted CD4⁺CD45RB^hi cells in 100 μL of PBS were transferred intravenously into Rag1^−/− mice with or without 10⁵ CD4⁺CD25^hi WT or A2BR^−/− Tregs through the retro-orbital venous sinus. Mice developed clinical signs of colitis 3.5–4.5 weeks post transfer. Mice were observed daily and weighed weekly. To assess the clinical status of the recipient mice, aggregate clinical scores were assigned as described previously (23) on the day of injection, weekly thereafter, and at time of sacrifice. For histologic scores, colon tissue sections were stained with hematoxylin and eosin (H&E) as well as Alcian blue and periodic acid–Schiff solution. Colitis severity was graded semiquantitatively from 0 to 4 in a blinded fashion.

Analysis of tumor antigen-specific effector CTL activity in vivo was performed as described previously (20). Briefly, SIY (SIYRYYGL) peptide-pulsed CFSE and OVA-I (SIINFEKL) peptide-pulsed CFSE low splenocytes were mixed at a ratio of 1:1, and a total of 2 × 10⁷ cells in 100 μL of PBS were injected intraperitoneally into recipient animals. Draining lymph nodes (DLN) and spleen were then harvested 24 hours after adoptive transfer, and CFSE fluorescence intensity was analyzed by flow cytometry. Gating on CFSE⁺ cells, killing was calculated as 1-[(% SIY peptide-pulsed cells in B16-SIY tumor–bearing mice/% OVA-I–peptide-pulsed cells in B16-SIY tumor–bearing mice)/(% SIY peptide-pulsed cells in tumor-free mice/% OVA-I–peptide-pulsed cells in tumor-free mice)] and expressed as a percentage.

CD45 cell enrichment from tumor infiltrates was initially conducted. Single-cell suspensions of B16-SIY tumors from WT or A2BR^−/− mice were stained with biotinylated anti-CD45, followed by streptavidin MACS beads, and sorted on an AutoMACS (Miltenyi Biotec). For the Treg suppression assay, the CD45⁺ infiltrating cells were further stained by anti-CD25 and anti-CD4 and sorted by BD Aria for CD4⁺CD25^hi Tregs. For the dendritic cell (DC)–mediated T-cell proliferation assay, the CD45⁺ infiltrating cells were further stained by anti-CD11b and anti-CD11c and sorted by BD Aria for CD11b⁺CD11c⁺ DCs. 2 × 10⁵ 2C splenocytes per well were plated into 96-well bottom plates in CR10 medium as described above in the presence of 1 μg/mL SIY peptides with or without serially diluted sorted Tregs or DCs for 3 days. For both assays above, wells were pulsed with 1 μCi of [³H]thymidine (PerkinElmer) for 8–10 hours and counted. 2C T-cell proliferation was defined as the mean CPM of the response of the antigen-stimulated cells divided by the mean of the response of cells cultured without antigen. Positive and negative controls were run on each plate.

BM-DCs were prepared as described previously (24). In brief, femurs and tibiae from 8- to 12-week-old mice were collected and flushed with sterile HBSS twice. The resulting BM cells were resuspended in R10F (RPMI1640 medium containing 10% heat-inactivated FBS, 2 mmol/L l-glutamine, 1 mmol/L sodium pyruvate, 50 U/mL penicillin, 50 mg/mL streptomycin) plus 50 mmol/L 2-ME, and 5 ng/mL GM-CSF. A total of 2–3 × 10⁶ cells per bacteriological culture plate were cultured for 10 days, feeding cells on days 3 and 8 by adding 10 mL fresh medium, and on day 6 by replacing half of the culture medium. Nonadherent cells were collected on day 10 and verified to be at least 85%–95% CD11b⁺/CD11c⁺/MHC-II⁺/F4/80⁻/Gr1⁻ by flow cytometry before use in experiments.

BM-DMs were prepared as described previously (3). In brief, BM cells obtained as described above ere cultured overnight in standard tissue culture plates in the presence of 10 ng/mL M-CSF. Nonadherent cells from this initial culture were then transferred to low-attachment six-well plates (Corning Life Sciences) in 4 ml R5F containing 30% L929 conditioned medium and 10 ng/mL M-CSF for 7 days, adding 1.5 mL fresh medium on days 3 and 5. Cells were verified to be at least 90%–98% CD11b⁺/CD11c⁻/F4/80⁺/Gr1⁻ by flow cytometry before use in experiments.

Data were analyzed by GraphPad Prism Software (Version 7.02). For datasets involving two groups Student t test, for datasets involving more than two groups one-way ANOVA and post hoc analysis and for samples involving more than one variable we performed two-way ANOVA and post hoc analysis. Significance is indicated as *, P < 0.05; **, P < 0.01; ***, P < 0.001 and ****, P < 0.0001.

Deletion or pharmacologic blockade of A2BR delays growth of syngeneic MB49 bladder carcinoma tumors in a hematopoietic cell–dependent manner (4). To extend the scope of these findings using multiple syngeneic tumor models, A2BR^−/− mice were injected subcutaneously with parental or ovalbumin expressing lung (LLC1 and LLC1-OVA), colon adenocarcinoma (MC38), and melanoma (B16-SIY) syngeneic tumors. All these tumor cell lines showed slower tumor growth in A2BR^−/− (KO) mice as compared with WT mice (Fig. 1A–D). To determine which A2BR⁺ (WT) host cell populations contributed to tumor protection, BM chimeras (WT mice receiving KO BM, i.e., KO>WT; KO mice receiving WT BM, i.e., WT>KO, KO mice receiving KO BM, i.e., KO>KO, and WT mice receiving WT BM, i.e., WT>WT) were used to ablate the expression of A2BR selectively in hematopoietic and nonhematopoietic cells. We found that A2BR deletion only on hematopoietic cells effectively limited the growth of LLC1 (Fig. 1E) or B16-SIY (Fig. 1F), indicating the predominant importance of A2BR expression on the hematopoietic compartment for tumor development.

We profiled immune infiltrates within the tumor microenvironment by flow cytometry. At 14 days postinoculation of B16-SIY tumors, there were no significant differences in the percentages of CD4⁺, Treg (CD4⁺Foxp3⁺), NK (CD49b⁺NK1.1⁺), CD11b⁺CD11c⁺ cells, or granulocytic myeloid-derived suppressor cells (Gr1⁺CD11b⁺) in the tumor infiltrates of B16-SIY tumors in A2BR^−/− versus WT mice (Fig. 2A; Supplementary Fig. S2). In contrast, the frequencies of tumor-infiltrating total and tumor antigen–specific CD8⁺ T cells (Fig. 2A and B, respectively) increased in A2BR^−/− mice as compared with WT mice. The frequency of infiltration of IFNγ-secreting CD8⁺ T cells was also increased in tumors isolated from A2BR^−/− hosts (Fig. 2C). To test whether these phenotypic differences translated into a functional activity, we performed an in vivo cytotoxicity assay by transferring peptide pulsed splenocytes into WT or A2BR^−/− hosts bearing B16-SIY tumors as depicted in Fig. 2D. Of note, the ability to kill antigen-pulsed target cells was significantly improved in DLNs of B16-SIY–bearing A2BR^−/− mice compared with WT mice (Fig. 2E). Our data demonstrated that host A2BR deficiency led to increased tumor antigen–specific effector CD8⁺ T-cell infiltration into the tumor microenvironment, contributing to delayed tumor growth.

To evaluate the role of intrinsic A2BR expression on CD8⁺ T-cell immunity, equal numbers of CFSE-labeled WT CD90.1⁺CD8⁺ T cells and A2BR^−/− CD90.2⁺ CD8⁺ T cells were cotransferred into CD45.1 transgenic mice one day before B16F10 or B16-SIY inoculation. Proliferation and IFNγ secretion of transferred splenic A2BR^−/− CD8⁺ T cells were comparable with those of cotransferred WT CD8⁺ T cells (Supplementary Fig. S3A and S3B). Adoptive transfer of A2BR^−/− CD8⁺ T cells caused tumor inhibition in Rag1^−/− mice similar to inhibition caused by WT CD8⁺ T cells. These results suggested an insignificant role of CD8⁺ T-cell–intrinsic A2BR expression for antitumor immunity (Supplementary Fig. S3C).

To determine whether A2BR expression influenced tumor-associated Treg function, we performed an in vitro suppression assay and an in vivo protection from colitis assay using A2BR^−/− versus WT Tregs isolated from B16-SIY tumors or spleen, respectively. There were no significant differences in suppressive functions between A2BR^−/− and WT CD4⁺Foxp3⁺ Treg from B16-SIY–bearing mice (Supplementary Fig. S2). In line with this, WT and A2BR^−/− Tregs showed comparable ability to protect mice from colitis (Supplementary Fig. S4).

To dissect the contribution of A2BR expression in APCs on antigen-specific T-cell priming, CFSE-labeled naïve CD8⁺ T cells were transferred by intravenous injection into WT (A2BR^+/+) or A2BR^−/− mice prior to B16-SIY tumor challenge. Enhanced proliferation (CFSE^dim) of transferred splenic CD8⁺ T cells was found in A2BR^−/− hosts compared with WT hosts (Fig. 3A and B). There was also higher expression of activation markers (CD69 and CD44) and IFNγ production in transferred CD8⁺ T cells from spleens collected from B16-SIY–treated A2BR^−/− mice compared with that of WT mice (Fig. 3A and B). These results suggested that A2BR-deficient APCs enhanced priming and activation of tumor antigen–specific A2BR-proficient CD8⁺ T cells. More specifically, we observed that tumor-infiltrating myeloid APCs from A2BR^−/− mice displayed elevated activation and maturation markers (CD86 and MHC II expression) as compared with that of WT mice (Fig. 3C). Sorted A2BR^−/− infiltrating myeloid APCs had enhanced capacity to drive antigen-specific T-cell proliferation in a dose-dependent manner compared with that of WT APCs ex vivo (Fig. 3D). These data demonstrated that A2BR deficiency allowed tumor-infiltrating myeloid APCs to more competently facilitate expansion and activation of tumor antigen–specific CD8⁺ T cells.

We found that only the selective A2BR antagonist, PSB-603, blocked the suppression of IL12 from DCs stimulated with the nonselective adenosine agonist NECA and LPS (Fig. 3E). Conversely, IL12 secreted from myeloid DCs derived from BM were remarkably reduced by the treatment with the A2BR partial agonist, BAY-60-6583, but not by selective agonists for A1R, A2AR, or A3R (Fig. 3F). We found that NECA was only capable of reducing an LPS-induced TNFα⁺IL10⁻ subpopulation of BM-DMs and increasing a TNFα⁻IL10⁻ DC subpopulation when BM-DMs are proficient for A2BRs (Supplementary Fig. S5A). Compared with either WT or A2AR^−/− BM-DM, A2BR^−/− BM-DMs produced more TNFα and less of IL10 when stimulated with LPS (Supplementary Fig. S5B). Collectively, these results suggested that A2BR expression was important for polarization of myeloid APCs from tumor suppressor to tumor-promoting phenotypes in vivo and ex vivo.

To investigate the cell-intrinsic role of A2BRs in hematopoietic cell differentiation in tumor-bearing mice, Rag1^−/− hosts were reconstituted with equal numbers of WT and A2BR^−/− BM cells. The proportion of A2BR^−/− myeloid APCs, CD11bdimMHCII⁺ DCs and Gr-1⁺ cells, but not T cells, in the tumor microenvironment increased compared with WT cells through 7–21 days posttumor inoculation (Fig. 4A). There was a slight but significant increase in the proportion of A2BR^−/− NK cells (Fig. 4A). Consistent with the results shown in Fig. 3C, there was significantly higher expression of CD86 (Fig. 4B) and MHCII (Fig. 4C) in A2BR^−/− infiltrating myeloid APCs than WT mDC on days 14 and 21. In contrast, the general activation status of both splenic and tumor-infiltrating CD4⁺ and CD8⁺ T cells was similar between A2BR^−/− and WT as measured by CD69, CXCR3 expression and IFNγ accumulation upon restimulation (Supplementary Fig. S6A and S6B). We did not observe significant changes in activation markers such as CD44, CD69, CXCR3, CD11b, or CD107a in A2BR^−/− NK cells (Supplementary Fig. S7). These results implicate a specific role for cell-intrinsic A2BR signaling in the differentiation and activation of myeloid cells within tumors.

To confirm cell-intrinsic effects of myeloid A2BR ablation on tumor progression, LysMCre^+/−Adora2b^f/f mice were generated for myeloid-selective A2BR deletion. Myeloid-selective deletion of A2BR inhibited MB49 tumor growth (Fig. 4D) as compared with that of LysMCre^−/− littermate control mice. Likewise, deletion of A2BR on myeloid cells dramatically reduced lung metastases of B16F10 melanomas detected by luciferase activity and lung weight measurement (Fig. 4E–H). We next performed a detailed immuno-profiling of myeloid cells and DC populations using B16-SIY tumors. We observed that A2BR deletion particularly caused increased infiltration among the tumor-associated myeloid DCs and CD103⁺ DCs (Fig. 5A). To test whether the role of DCs in tumor growth and dissemination we crossed Adora2b^f/f mice with ItgaxCre^+/− mice (CD11cCre). Deletion of A2BRs from DCs delayed growth B16F10-luciferase cells but failed to delay decrease lung dissemination (Fig. 5B–D) suggesting A2BR expression among DCs promoted tumor growth while A2BR expression of monocytes and/or Gr1⁺ cells promoted lung dissemination.

Cell-based therapies are efficacious for treating leukemia but have failed to slow the growth solid tumors, possibly due to the suppressive tumor microenvironment (25). High adenosine in the TME can suppress tumor rejection by the immune system (1). Therefore, to extend our findings to a more clinically relevant setting, we evaluated the antitumor effect of pharmacologic A2BR blockade on adoptive T-cell therapy. In line with previously published results (20), transfer of SIY/anti-CD28 and IL2 prestimulated SIY-specific 2C T cells alone failed to control the growth of B16-SIY tumors (Fig. 6). Consistent with a role for host A2BRs in promoting tumor growth, treatment with the selective A2BR antagonist MRS-1754 caused tumor growth inhibition in both B16-SIY (Fig. 6) and MC38 (Supplementary Fig. S8) tumor models as a monotherapy. Moreover, combining daily MRS1754 treatment with adoptive 2C T-cell therapy achieved synergistic antitumor efficacy (Fig. 6). Overall, our study suggested that APC expression of A2BR in the tumor microenvironment inhibits antigen-specific antitumor immune responses, thus can be exploited as a way to improve current strategies of cancer immunotherapy, including cell-based therapies.

Adenosine A2B receptors are expressed in tumor cells and immune cells (4, 7, 11). Blockade or pharmacological deletion of A2BRs causes activation of both tumor-associated T cells and APCs in vivo (4). However, it was not clear which immune cell subtype have intrinsic A2BRs. Also, although tumor expression of A2BR is important for lung colonization of tumors (6, 7), this finding does not rule out a role for A2BRs on immune cells in contributing to this effect. Here, we provided evidence that cell-intrinsic signaling by A2BRs on APCs is involved in adenosine-mediated suppression of antitumor T-cell responses and for enhancing dissemination of tumors to the lung.

Activation of macrophages and DCs differentiated from bone marrow cells can be suppressed by A2BR signaling (10, 11). A2BR signaling in DCs strongly suppressed activation of T cells in vitro (11). Accordingly, in vivo models of inflammatory and infectious disease models suggest that A2BR signaling can suppress macrophage activation and phagocytosis, potentially leading to increased bacterial pathogenesis but reduced sepsis (16, 26), suggesting a role for myeloid cell A2BRs as a tumor-promoting mechanism. Deletion of A2BR or pharmacologic targeting slows the growth of bladder carcinoma in a T-cell–dependent manner (4). However, it was not clear if myeloid cells and DCs intrinsically played a major role in producing this effect and whether similar mechanisms are at play in other tumors. Here, we addressed this question by extending the scope of previous observations showing antitumor effects of A2BR deletion in multiple syngeneic tumor models and defining the hematopoietic compartment and specific APCs as cellular targets of A2BRs in the tumor microenvironment. A2BR expression is mostly observed in myeloid cells and myeloid DCs in PBMCs (ref. 27; https://www.proteinatlas.org/). Human DCs are susceptible to adenosine suppression ex vivo (28). Further investigation is required using human tumor samples to find whether increased expression of A2BR particularly in tumor versus tumor-infiltrating APCs is associated with poor survival among these patients.

A2BR signaling promotes tumor metastasis (6, 7, 29, 30). A2BR blockade inhibited both spontaneous metastasis of primary tumors and lung colonization of tumors after systemic delivery. Mechanistically, A2BR expression in tumor cells decreases cell-to-cell contact and increases tumor cell survival through the ERK pathway (7, 8). Breast tumor cells express intrinsic A2B receptors that facilitate metastasis (6, 7). A2BR blockade or silencing CD73⁺ in such tumors does not influence T cells or NKs (7). Our study suggests that A2BR expression in monocytes and/or Gr1⁺ myeloid cells but not DCs could also promote tumor colonization, especially at a later time point (week 2) after systemic delivery of the tumor cells; these data emphasized the importance of these subpopulations in regulation of tumor metastasis (31–33). One key difference between these studies and our study is the use of CD73⁺ cells for tumor cell–intrinsic effects (6, 7, 30), and the use of A2BR^low, CD73^low B16F10 cells to uncover antitumor immune effects, respectively. Therefore, depending on the model, immune versus tumor-related effects of A2BR on metastasis may vary. Therefore, A2BRs on both tumor cells and immune cells, particularly myeloid cells, may be responsible for tumor invasiveness with varying degrees of influence depending on the CD73 status of the tumor cell. This concept requires additional investigation.

Adenosine targets both A2ARs and A2BRs to suppress immune responses (3–5, 20, 34, 35). Myeloid expression of adenosine A2ARs suppresses antitumoral T and NK-cell responses (3). Also, growth of certain syngeneic tumors such as MCA205 sarcomas or immunogenic CL8-1 melanomas are more sensitive to A2AR deletion than A2BR deletion (5, 36). Both A2ARs and A2BRs couple to Gs and increase cAMP to suppress immune responses (1). A2BRs also couples to Gq G proteins and has cAMP-independent effects. A2ARs are widely distributed on immune cells (2), whereas A2BRs more preferentially expressed on myeloid immune cells (9). Therefore, A2AR and A2BR signaling can cause both overlapping and nonoverlapping effects in regulation of immune cell function causing differential effects in different tumor microenvironments. Another potential difference may be due to the affinities of these receptors for adenosine. A2ARs have higher affinity, and deletion of A2ARs can play important roles in homeostasis of immune cells systemically, while low-affinity A2BR signaling is preferentially involved in local pathologic reactions associated with high adenosine generation (37, 38). Therefore, tumors with high adenosine accumulation may be more sensitive to A2BR blockade. Another possibility is based on the role of A2ARs in regulating the expression of A2BRs, suggesting that deletion of A2ARs may also reduce A2BR signaling indirectly (39). Future studies are needed to test if deletion or targeting both receptors especially in myeloid cells will have a synergistic or additive effect on tumor growth and dissemination.

In summary, our study identified cell-intrinsic A2BR expression on APCs as a suppressor of antitumor immune responses. There are distinct subtypes of APCs such as CD103⁺ DCs that are mainly responsible for antigen-cross presentation to CD8⁺ T cells and myeloid DCs that activate CD4⁺ T cells and produce tolerogenic versus immunostimulatory cytokines such as IL10 versus IL12, respectively, to other immune cells including CD8⁺ T cells depending on the microenvironment (40). Additional studies are needed to further dissect the roles of A2BRs in DC subtypes. Overall, our findings suggested that cell-based therapies such as adoptive cell therapy or DC vaccines, and immunotherapies can become more efficacious in combination A2BR inhibition.

No potential conflicts of interest were disclosed.

Conception and design: S. Chen, B. Zhang, C. Cekic

Development of methodology: S. Chen, I. Akdemir, J. Fan, B. Zhang, C. Cekic

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): S. Chen, J. Fan, J. Linden, B. Zhang, C. Cekic

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): S. Chen, I. Akdemir, B. Zhang, C. Cekic

Writing, review, and/or revision of the manuscript: S. Chen, J. Linden, B. Zhang, C. Cekic

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): S. Chen, I. Akdemir, B. Zhang, C. Cekic

Study supervision: B. Zhang, C. Cekic

The authors thank Long Wang and Lishi Sun (University of Texas Health Science Center at San Antonio) for their technical support with the in vivo mouse studies. The authors also thank Ali Can Savas, Merve Kayhan, and Altay Koyas (Bilkent University, Ankara, Turkey) for their technical assistance with the in vitro studies. This research was supported in part by NIH grant CA149669, a Melanoma Research Alliance Pilot Award (to B. Zhang), Northwestern University Robert H. Lurie Comprehensive Cancer Center Flow Cytometry Facility, and a Cancer Center Support Grant (NCI CA060553). This work is also supported in part by European Molecular Biology Organization (EMBO) Installation Grant 3297 and by 1001 - Scientific and Technological Research Projects Funding Program grant (215S729) from Scientific and Technological Research Council of Turkey (TUBITAK; to C. Cekic).

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