PD-L1 ag T omot self-oleranc es macrophag fect T er€¦ · l, CD4+ T cells in day 21 orthotopic...

Articleshttps://doi.org/10.1038/s41590-020-0620-x

1S. Arthur Localio Laboratory, Department of Surgery, New York University School of Medicine, New York, NY, USA. 2Department of Radiation Oncology, New York University School of Medicine, New York, NY, USA. 3Department of Pathology, New York University School of Medicine, New York, NY, USA. 4Department of Cell Biology, New York University School of Medicine, New York, NY, USA. 5Department of Medicine, New York University School of Medicine, New York, NY, USA. ✉e-mail: [email protected]

PD-L1, the cognate ligand for PD-1, is a 40 kDa transmembrane protein with a short cytoplasmic tail1. The PD-L1–PD-1 axis has become a central target of immunotherapy approaches

in the clinic2. PD-L1 is robustly upregulated on tumor cells and on intra-tumoral macrophages and dendritic cells3. However, the sig-nificance of T cell expression of PD-L1 is less certain.

Ligation of PD-L1 in human T cells was previously shown to induce IL-10 expression and promote cellular apoptosis4. As such, PD-L1 autoantibodies have been implicated in the progression of rheumatoid arthritis by the induction of aberrant T cell responses4. It was reported previously that peak expression of PD-L1 on CD8+ T cells occurred during the contraction phase of an immune response and that PD-L1 blockade at this stage reduced the num-bers of effector CD8+ T cells5. PD-L1 was needed to maintain the levels of Bcl-xL in activated CD8+ T cells. Notably, the reported expression of PD-L1 by tumor-infiltrating lymphocytes (TILs) and the demonstration that PD-L1hi CD8+ TILs express more IFN-γ and CD107a than do PD-L1lo CD8+ TILs, suggested that PD-L1hi TILs are functional effector cells6. However, engagement of PD-L1 induced T cell apoptosis and resulted in loss of therapeutic effects. Nevertheless, it remains uncertain whether PD-L1 ligation in T cells is an important mechanism of tumor immune evasion.

We found that PD-L1 is upregulated on T cells in cancer in response to antigen-presentation and as a consequence of sterile

inflammatory cues. Moreover, PD-L1 ligation in T cells induces intracellular signaling that is equally suppressive to that of PD-1. PD-L1+ T cells also promote STAT6-dependent M2-like macro-phage differentiation and suppress neighboring effector T cells via the canonical PD-L1–PD-1 axis. Collectively, our data suggest that T cell expression of PD-L1 maintains intra-tumoral immune tolerance via fate-determining ‘back-signaling’ and promotion of a suppressive phenotype in the adjacent innate and adaptive immune compartments.

ResultsT cell expression of PD-L1 in cancer is regulated by antigen presentation, JAK-STAT signaling, and soluble inflammatory mediators. We investigated the prevalence of PD-L1 expression on T cells in a preinvasive autochthonous model of pancreatic ductal adenocarcinoma (PDA) using p48-cre; KrasG12D/+ (KC) mice, which express oncogenic Kras in their pancreatic progenitor cells, in an invasive orthotopic PDA model using tumor cells derived from Pdx1-cre; KrasG12D/+; Tp53R172H/+ (KPC) mice, which express mutant Kras and p53, in a KPC liver metastasis model, and in human dis-ease. Approximately 50% of T cells expressed PD-L1 in pancreata of 6-month-old KC mice (Fig. 1a). In orthotopic KPC tumors, PD-L1 was expressed in ~40% of CD4+ T cells and ~60% of CD8+ T cells, compared to minimal expression in T cells of normal pancreas

PD-L1 engagement on T cells promotes self-tolerance and suppression of neighboring macrophages and effector T cells in cancerBrian Diskin1, Salma Adam1, Marcelo F. Cassini1, Gustavo Sanchez1, Miguel Liria1, Berk Aykut1, Chandan Buttar1, Eric Li1, Belen Sundberg1, Ruben D. Salas1, Ruonan Chen1, Junjie Wang1, Mirhee Kim1, Mohammad Saad Farooq1, Susanna Nguy2, Carmine Fedele3, Kwan Ho Tang3, Ting Chen 3, Wei Wang1, Mautin Hundeyin1, Juan A. Kochen Rossi1, Emma Kurz1, Muhammad Israr Ul Haq1, Jason Karlen1, Emma Kruger1, Zennur Sekendiz1, Dongling Wu1, Sorin A. A. Shadaloey1, Gillian Baptiste1, Gregor Werba1, Shanmugapriya Selvaraj3, Cynthia Loomis3,4, Kwok-Kin Wong5, Joshua Leinwand1 and George Miller 1,4 ✉

Programmed cell death protein 1 (PD-1) ligation delimits immunogenic responses in T cells. However, the consequences of programmed cell death 1 ligand 1 (PD-L1) ligation in T cells are uncertain. We found that T cell expression of PD-L1 in cancer was regulated by tumor antigen and sterile inflammatory cues. PD-L1+ T cells exerted tumor-promoting tolerance via three distinct mechanisms: (1) binding of PD-L1 induced STAT3-dependent ‘back-signaling’ in CD4+ T cells, which prevented activa-tion, reduced TH1-polarization and directed TH17-differentiation. PD-L1 signaling also induced an anergic T-bet−IFN-γ− pheno-type in CD8+ T cells and was equally suppressive compared to PD-1 signaling; (2) PD-L1+ T cells restrained effector T cells via the canonical PD-L1–PD-1 axis and were sufficient to accelerate tumorigenesis, even in the absence of endogenous PD-L1; (3) PD-L1+ T cells engaged PD-1+ macrophages, inducing an alternative M2-like program, which had crippling effects on adaptive antitumor immunity. Collectively, we demonstrate that PD-L1+ T cells have diverse tolerogenic effects on tumor immunity.

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(Fig. 1b). T cell expression of PD-L1 was also higher in tumor than in spleen (Fig. 1b and Supplementary Fig. 1a). Immune fluorescence microscopy confirmed the selective T cell expression of PD-L1 in PDA (Fig. 1c). By contrast, PD-L2 was minimally expressed on tumor-infiltrating T cells (Fig. 1d). T cell expression of PD-L1 increased with progressive oncogenesis (Fig. 1e). Notably, PD-L1 was mildly upregulated in T cells in chronic pancreatitis, but to a far lesser extent than in PDA (Fig. 1f). In PDA liver metastases, tumor-infiltrating CD4+ and CD8+ T cells also upregulated PD-L1, albeit to lower levels than in primary tumor (Fig. 1g). In human PDA, PD-L1 expression was highly upregulated in T cells infiltrating tumor tissues and was mildly elevated in T cells in peripheral blood mononuclear cells (PBMC) of patients with cancer compared to PBMC of healthy volunteers (Fig. 1h). Notably, expression of PD-L1 in T cells in PDA was higher than its expression in tumor cells or in B cells and macrophages (Fig. 1i). Overall, despite their relative scarcity in the tissue microenvironment (TME)7, T cells constituted ~30% of PD-L1+ cells in PDA (Fig. 1j). Analysis of other cancer sub-types suggested that expression of PD-L1 was highly upregulated on intra-tumoral T cells in murine melanoma but was expressed at lower levels in T cells in colorectal cancer (Fig. 1k). Comparison of the phenotype of PD-L1+ T cells to PD-L1− T cells in PDA sug-gested that PD-L1 is highly coexpressed with surface activation markers, diverse co-stimulatory and checkpoint receptors, TH1-, TH2-, and Treg-family cytokines, and transcription factors in both CD4+ and CD8+ T cells (Supplementary Fig. 1b–g). By contrast, PD-L1− T cells exhibited a comparatively non-activated phenotype. In particular, 4-1BB and LAG-3 were almost exclusively expressed in PD-L1+ T cells in the PDA TME (Fig. 1l).

To determine whether antigen presentation drives T cell expres-sion of PD-L1 in vivo, we analyzed PD-L1 expression in Ova-pentamer+ and Ova-pentamer− CD8+ T cells infiltrating KPC tumors engineered to express ovalbumin (Ova)8. Intra-tumoral anti-gen-specific T cells expressed markedly higher PD-L1 compared to non-antigen-restricted T cells (Fig. 2a). Antigen presentation simi-larly upregulated PD-L1 expression in vitro in Ova-restricted CD4+ T cells (Fig. 2b). Antibody-based ligation of the CD3/CD28 co-receptors in T cells also progressively upregulated PD-L1 in mouse and human systems (Fig. 2c,d). Mechanistically, induction of PD-L1 expression in T cells by antigen presentation or co-receptor ligation required JAK-STAT signaling (Fig. 2e,f). To investigate whether sterile inflammatory mediators in the PDA TME can similarly modulate PD-L1 expression on T cells, we induced Ova-restricted CD8+ T cells to express PD-L1 by cross-presentation of antigen and simultaneously blocked selected cytokine or chemokine sig-naling networks that are either known to be upregulated in PDA

(IL-6, IL-4, IL-17 and IL-10) or are purported to regulate PD-L1 expression in APC (IFN-γ and IL-27) (refs. 9–13). Blockade of IL-4, IFN-γ and IL-27 partially mitigated PD-L1 expression in Ova-restricted T cells (Fig. 2g). We confirmed that T cells that infiltrate PDA tumors in interferon gamma-deficient (Ifng−/−) and interleu-kin 27 receptor-deficient (Il27r−/−) hosts expressed lower PD-L1 (Fig. 2h). Collectively, these data suggest that PD-L1 expression on T cells is modulated by presentation of tumor antigen and inflam-matory cues. To determine whether PDA tumor cells directly induce PD-L1 expression in T cells, we cocultured splenic T cells with KPC cells; however, tumor cells or their conditioned media failed to upregulate PD-L1 in T cells (Fig. 2i).

PD-L1 expression does not affect resting T cell phenotype but mit-igates T cell activation. Splenic T cells (Supplementary Fig. 2a,b) and their thymic precursors (Supplementary Fig. 2c–f) exhibited similar preactivation phenotypes in wild-type (Pdl1+/+) and PD-L1-deficient (Pdl1−/−) mice. However, PD-L1 deletion enabled T cells to achieve higher activation states after CD3/CD28 coligation, as Pdl1−/− CD8+ T cells adopted an enhanced cytotoxic phenotype (Supplementary Fig. 2g), and Pdl1−/− CD4+ T cells expressed elevated TH1-, TH2-, and Treg-associated transcription factors and cytokines (Supplementary Fig. 2h). Collectively, these data suggest that whereas Pdl1+/+ and Pdl1−/− T cells have similar preactivation states, PD-L1 expression in T cells suppresses cellular activation.

Conditional deletion of PD-L1 in T cells enhances adaptive tumor immunity and activates tumor-associated macrophages. To deter-mine the effect of T cell expression of PD-L1 on tumor immunity, we conditionally deleted PD-L1 in T cells using Cd4-cre; Pdl1fl/fl mice (Supplementary Fig. 2i) and challenged these mice, along with lit-termate controls, with orthotopic PDA tumors. Cd4-cre; Pdl1fl/fl mice developed smaller tumors than did littermate controls (Fig. 3a). Moreover, we noted marked expansion and activation of tumor-infiltrating T cells in Cd4-cre; Pdl1fl/fl mice (Fig. 3b,c). Analysis of the inflammatory TME by single-cell RNA sequencing (scRNA-seq) revealed that tumor-infiltrating T cells in Cd4-cre; Pdl1fl/fl hosts upreg-ulated expression of activating signal transducers (for example Zap70, Bcl3, Tyk2, and Syk), inflammatory mediators and activation mark-ers (Il6ra, Ccr7, Pdcd4 and Cd83), and fate-determining transcription factors (Tcf7 and Nfx1) (Fig. 3d–f). Consistent with these data, assess-ment of gene ontology (GO) revealed increased T cell activation and differentiation and upregulated protein kinase signaling in tumor-infiltrating T cells in Cd4-cre; Pdl1fl/fl mice (Fig. 3g). In addition to activating T cells, tumor-associated macrophages (TAMs) in Cd4-cre; Pdl1fl/fl hosts exhibited an immunogenic phenotype. For example, the

Fig. 1 | T cells are a primary source of PD-L1 in PDA. a, CD3+ T cells from pancreata and spleens of 6-month-old KC mice were gated and tested for expression of PD-L1 by flow cytometry. b, Pdl1+/+ mice bearing orthotopic KPC tumors were killed on day 21. Tumors and spleens were harvested and CD4+ and CD8+ T cells were gated by flow cytometry and tested for expression of PD-L1. T cells from normal pancreata were also tested. Representative contour plots and quantitative data are shown. Gates are based on isotype control. c, Paraffin sections of human PDA tumors were stained using antibodies specific for PD-L1, CD3 and CK19. Nuclei were stained using 4,6-diamidino-2-phenylindole (DAPi). Representative image is shown (scale bar, 10µm). d, Comparison of PD-L1 and PD-L2 expression in CD4+ and CD8+ T cells infiltrating day 21 orthotopic KPC tumors by flow cytometry. e, Pdl1+/+ mice were orthotopically administered KPC-derived tumor cells. Cohorts were killed on days 7, 14 or 21 and PDA-infiltrating CD4+ T cells were tested for expression of PD-L1. Representative contour plots, including day 7 isotype control, and quantitative data are shown. f, Comparison of PD-L1 expression in CD4+ T cells in normal pancreata, chronic pancreatitis, and orthotopic PDA tumors. g, Comparison of PD-L1 expression in CD4+ and CD8+ T cells in KPC liver metastases versus orthotopic KPC tumors. h, TCRαβ+ T cells in PBMC of healthy persons and in PMBC and tumor of patients with PDA were tested for expression of PD-L1. Representative histograms and quantitative data are shown. Gates are based on isotype controls. i, T cells, tumor cells, B cells, NK cells and macrophages from day 21 orthotopic KPC tumors were analyzed for expression of PD-L1 by flow cytometry. j, KPC tumors were harvested on day 21 and single-cell suspensions were made. Live cells were gated by flow cytometry and were analyzed for PD-L1 expression. PD-L1+ and PD-L1− cells were subgated and were analyzed for coexpression of CD45 and CD3. Representative contour plots are shown. k, CD3+ T cells from day 21 B16 melanoma and MCA38 colon cancer tumors were analyzed for PD-L1 expression. Representative contour plots compared to isotype control and quantitative data are shown. l, CD4+ T cells in day 21 orthotopic KPC tumors were tested for coexpression of PD-L1 with 4-1BB and LAG-3. Representative contour plots and quantitative data comparing 4-1BB and LAG-3 expression on PD-L1+ and PD-L1− cells are shown. Each experiment was repeated at least three times with similar results (*P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001). PE, phycoerythrin.

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CD68lo macrophage cluster exhibited upregulated expression of extra-cellular receptors (Timp2, B2m, Cd74), cell surface antigens (H2-K1), and select chemokines (Ccl6, C1qa, Mif) (Fig. 3h). Further, ingenuity pathway analysis revealed that TAMs in tumors of in Cd4-cre; Pdl1fl/fl

mice increased IL-8, CXCR4, mTOR, and EIF2 signaling (Fig. 3i). Analysis of upstream regulators indicated upregulated IL-6 signaling and diminished IL-10 signaling in TAMs in tumors of Cd4-cre; Pdl1fl/fl hosts (Fig. 3j,k). Accordingly, we found enrichment for GO terms

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associated with positive regulation of the adaptive immune response in tumor-infiltrating macrophages in Cd4-cre; Pdl1fl/fl mice (Fig. 3l). Collectively, these data suggest that conditional deletion of PD-L1 in T cells is tumor-protective and activates intra-tumoral T cells and macrophages.

Engagement of PD-L1 by PD-1 induces inhibitory signaling in T cells and drives TH17 differentiation. As PD-L1 deletion in T cells is associated with their enhanced capacity for activation, we postulated that ligation of PD-L1 induces suppressive back-sig-naling in T cells independent of the canonical PD-L1–PD-1 axis.

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Fig. 2 | Regulation of PD-L1 expression in T cells. a, Pdl1+/+ mice were implanted with orthotopic KPC tumor cells engineered to express ovalbumin. PDA tumors were harvested on day 21 and CD8+ T cells were gated and tested for both Ova-pentamer staining and PD-L1 expression. PD-L1 expression on Ova-pentamer+ cells and Ova-pentamer− cells was analyzed. This experiment was repeated three times with similar results. b, Ova-restricted CD4+ T cells from OT-ii mice were stimulated in vitro with Ova323–339 peptide or vehicle and were tested for PD-L1 expression at 72 h. Representative contour plots and quantitative data are shown. This experiment was repeated twice. c, Polyclonal CD4+ and CD8+ T cells from Pdl1+/+ mouse spleen or human PBMC were stimulated in vitro by CD3/CD28 coligation or were mock stimulated and tested for PD-L1 expression at 72 h. Representative contour plots and quantitative data are shown. This experiment was repeated more than three times with similar results. d, Polyclonal CD4+ and CD8+ T cells from Pdl1+/+ or Pdl1−/− mice were stimulated in vitro by CD3/CD28 coligation and were tested for PD-L1 expression at 24 h, 48 h and 72 h. This experiment was repeated twice. e, Polyclonal CD4+ and CD8+ T cells from Pdl1+/+ mice were stimulated in vitro by CD3/CD28 coligation and were tested for PD-L1 expression at 72 h. Selected wells were treated with JAK-STAT, mTOR, AKT or Pi3K inhibitors. This experiment was repeated three times with similar results. f, Ova-restricted CD4+ T cells from OT-ii mice were stimulated in vitro with Ova323–339 peptide or vehicle and were tested for PD-L1 expression at 72 h. Selected wells were treated with JAK-STAT, mTOR, AKT or Pi3K inhibitors. This experiment was repeated three times with similar results. g, OT-i T cells were tested for PD-L1 expression at 72 h after presentation of Ova257–264 peptide. Select wells were treated with antibodies targeting specific inflammatory mediators or receptors. This experiment was repeated four times with similar results. h, CD4+ and CD8+ T cells infiltrating day 21 orthotopic KPC tumors in Pdl1+/+, Ifng−/− and Il27r−/− hosts were tested for expression of PD-L1. This experiment was repeated twice with similar results. i, Polyclonal splenic T cells from Pdl1+/+ mice were activated by CD3/CD28 coligation and were selectively cocultured with KPC tumor cells (1:1 ratio) or with tumor-cell-conditioned medium before assessment of PD-L1 expression. This experiment was repeated three times with similar results (*P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001). MFi, mean fluorescence intensity.




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Fig. 3 | Conditional deletion of PD-L1 in T cells enhances adaptive tumor immunity and activates tumor-associated macrophages. a–c, Cd4-cre; Pdl1fl/fl mice and littermate controls were administered orthotopic KPC tumors and were killed at three weeks. Tumor weight (a), CD3+ T cell infiltrate (b), and T cell expression of iFNγ and TNF (c) were measured. d–l, Cd4-cre; Pdl1fl/fl mice and littermate controls were administered orthotopic KPC tumors and were killed at three weeks. scRNA-seq was performed on FACS-purified CD45+ tumor-infiltrating leukocytes. d, The distribution of cellular clusters was determined using the t-SNE algorithm. Each cluster is identified by a distinct color. Bar graph (e) and violin plots (f) comparing normalized (norm.) log expression of genes in the T cell cluster for Cd4-cre; Pdl1fl/fl (n = 1) and control (n = 2) groups. g, Network analysis shows GO relationship for leukocyte differentiation pathways and stress-activated protein kinase signaling in tumor-infiltrating Cd4-cre; Pdl1fl/fl T cells compared to control. h, Violin plots comparing normalized log expression of selected genes in the CD68lo macrophage cluster. i, ingenuity pathway analysis (iPA)-based canonical pathway perturbations in the CD68lo macrophage cluster for the Cd4-cre; Pdl1fl/fl versus control T cell groups. j–k, Bubble plots depicting regulation of iL-6 (j) and iL-10 (k) signaling in CD68lo macrophages in tumors of Cd4-cre; Pdl1fl/fl mice compared to controls. l, Network analysis shows GO relationship for adaptive immune response in tumor-associated CD68lo macrophages in Cd4-cre; Pdl1fl/fl mice compared to controls (*P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001). MΦ, macrophage.




To test this, we activated Ova-restricted CD8+ T cells by presentation of Ova257-264 peptide in the presence of PD-1 Fc or IgG Fc control. PD-1 Fc prevented CD8+ T cell priming as evidenced by reduced expression of surface activation markers, immunogenic cytokines, and transcription factors (Fig. 4a–g). In a similar manner, ligation of PD-L1 abrogated antigen-restricted CD4+ T cell activation (Fig. 4h–l). PD-L1 ligation also mitigated αCD3/αCD28-mediated activation of CD4+ (Supplementary Fig. 3a,b) and CD8+ (Supplementary Fig. 3c,d) T cells and inhibited T cell proliferation (Supplementary Fig. 3e). We observed a dose-response effect of PD-1 Fc treat-ment in Pdl1+/+ T cells (Supplementary Fig. 3f–l). PD-1 Fc did not affect T cell activation in Pdl1−/− T cells, which confirms that the specificity of effects is restricted to PD-L1 ligation (Supplementary Fig. 3m–o). To validate our findings, we transduced Pdl1−/− T cells with PD-L1 using a lentiviral vector (Supplementary Fig. 4a,b). PD-1 Fc treatment induced a suppressive phenotype in the Pdl1−/− T cells that were transduced with a PD-L1-expressing vector (Supplementary Fig. 4c–f) but not in T cells transduced with the empty vector (not shown). Ligation of PD-L1 also mitigated the activation of human PBMC-derived T cells (Fig. 4m). Notably, the suppressive effects of PD-L1 signaling were comparable to that of PD-1 signaling in T cells (Supplementary Fig. 5a–c). Furthermore, similar inhibitory effects of PD-L1 engagement were observed in tumor-infiltrating PD-L1+ T cells (Supplementary Fig. 5d,e).

To elucidate the transcriptomic changes in T cells as a conse-quence of PD-L1 ligation, we performed RNA-seq on human T cells treated with PD-1 Fc or IgG Fc. PD-L1 ligation induced marked global transcriptomic changes in T cells (Fig. 4n and Supplementary Fig. 6a). Most notably, we observed downregulation of genes that are associated with the immunogenic response and IFN-γ signal-ing, including IRF7, MXI, IFI44L, OAS3, IFIT3 and CXCL13. GO analysis confirmed that engagement of PD-L1 downregulated Type I IFN signaling, T cell cytotoxicity, antigen processing and presenta-tion, and innate immune response (Fig. 4o). Accordingly, analysis of upstream regulators demonstrated that PD-L1 binding in T cells diminished the T cell antigen receptor signaling, IFN-γ, IFN-α, IFN-α receptor, and TNF associated pathways (Supplementary Fig. 6b–f). Ingenuity analysis indicated that canonical pathways that are downregulated by PD-L1 engagement include IFN sig-naling, TH1 and TH2 pathways, as well as CD28, OX40, and ICOS co-stimulation (Fig. 4p). Gene set enrichment analysis (GSEA) con-firmed that genes in the ‘antigen processing and presentation’ and ‘chemokine signaling pathway’ were markedly reduced upon PD-L1 ligation (Fig. 4q,r).

To determine the consequences of inhibitory PD-L1 signaling in situ in PDA, we treated patient-derived organotypic tumor spher-oids (PDOTS) with PD-1 Fc or IgG Fc using a microfluidic system that was recently validated as a model for immunotherapeutic testing14. Treatment of PDOTS with PD-1 Fc accelerated spheroid growth and suppressed intra-tumoral T cell phenotype compared

to treatment with IgG Fc (Supplementary Fig. 6g,h). In aggregate, these data indicate that PD-L1 ligation in T cells has tumor-promot-ing tolerogenic consequences in both mouse and human systems, whereas deletion of PD-L1 in the T cell compartment alone is suf-ficient to confer tumor immunity.

As our transcriptomic analyses suggested that PD-L1 ligation alters lymphocyte differentiation pathways, we directly studied the consequences of PD-L1 engagement on TH cellular polarization. Ligation of PD-L1 in CD4+ T cells reduced TH1 (Tbet and IFN-γ) and TH2 (GATA3) differentiation but promoted TH17 (Rorγt and IL-17F) programming after αCD3/αCD28 activation (Fig. 5a,b). Nevertheless, IL-17A was downregulated. Ligation of PD-L1 simi-larly increased IL-17F expression in CD8+ T cells (Supplementary Fig. 6i). The effect of PD-L1 ligation on mitigating TH1 and TH2 differentiation but promoting a partial TH17 phenotype was con-firmed by quantitative PRC (qPCR) (Fig. 5c,d). Similarly, treatment of OT-II T cells with PD-1 Fc in the context of Ova323-339 peptide presentation reduced CD4+ T cell expression of Tbet but upregu-lated Rorγt (Fig. 5e,f). FoxP3 expression was unchanged (Fig. 5g). The promotion of partial TH17 polarization and reduced TH1 dif-ferentiation as a consequence of PD-L1 ligation was corroborated in human T cells at both the protein (Fig. 5h) and RNA (Fig. 5i) levels. Accordingly, PD-L1 engagement upregulated STAT3 sig-naling in CD4+ T cells, associated with TH17 polarization, but reduced STAT1 signaling, which drives TH1 differentiation (Fig. 5j) (refs. 15,16). p38 MAP Kinase, PI3K–Akt, ITK and ERK signaling were also diminished after ligation of PD-L1, which is consistent with global suppression of T cell activation (Fig. 5j,k). SOCS3, which inhibits STAT3 activation, was similarly downregulated (Fig. 5k) (ref. 17). Upstream analysis of transcriptomic data cor-roborated that PD-L1 ligation enriched STAT3 signaling pathways in T cells but inhibited STAT1-associated pathways (Fig. 5l,m). Consistent with these observations, GSEA indicated that genes in the FoxO signaling pathway, which is upregulated by STAT3 signaling but inhibited by PI3K–Akt and MAP kinase, were increased as a consequence PD-1 Fc treatment (Fig. 5n) (refs.18–20). Immunoblotting confirmed that PD-L1 signaling reduced FoxO1 phosphorylation, which is indicative of increased FoxO signaling (Fig. 5k) (ref.18). Moreover, STAT3 inhibition (Fig. 5o,p) or dele-tion (Fig. 5q) abrogated the upregulation of Rorγt and IL-17F that is associated with PD-L1 ligation in T cells. Collectively, these data suggest that PD-L1 signaling in T cells promotes partial TH17 dif-ferentiation via STAT3 while mitigating TH1 and TH2 polarization.

PD-L1+ T cells suppress neighboring T cells in the TME. In addi-tion to inhibitory back-signaling, PD-L1-expressing T cells can theoretically suppress immunity via ligation of PD-1 on neighbor-ing T cells in the TME. Therefore, we investigated the immune-suppressive capacity of PD-L1+ T cells on neighboring T cells via the PD-L1–PD-1 axis. We stimulated Ova-restricted CD8+ T cells

Fig. 4 | Ligation of PD-L1 induces suppressive back-signaling in T cells. a–g, Antigen-restricted CD8+ OT-i T cells were stimulated for 72 h with Ova257–264 peptide in the presence of PD-1 Fc or control igG Fc. T cell activation was determined by expression of CD44 (a), LFA-1 (b), CD107a (c), TNF (d), iFNγ (e), Tbet (f), and EOMES (g). This experiment was repeated four times with similar results. h–l, Antigen-restricted CD4+ OT-ii T cells were stimulated for 72 h with Ova323–339 peptide in the presence of PD-1 Fc or igG Fc. T cell activation was determined by expression of CD44 (h), LFA-1 (i), CD69 (j), TNF (k), and Tbet (l). This experiment was repeated more than four times with similar results. m, Polyclonal human PBMC-derived T cells were stimulated for 72 h by CD3/CD28 coligation in the presence of PD-1 Fc or igG Fc. CD3+ T cell activation was determined by expression of CD69, CD62L, iCOS, TNF, Granzyme B and LFA-1. Selected representative histograms and quantitative data are shown. This experiment was repeated three times using PBMC from different volunteers with similar results. n–r, Polyclonal human PBMC-derived T cells were stimulated by CD3/CD28 coligation in the presence of PD-1 Fc or igG Fc. Cells were harvested at 72 h for analysis by RNA-seq in triplicate. n, Volcano plot depicting differentially expressed genes for PD-1 Fc treatment versus control. o, Top scoring GO terms related to innate inflammatory and immune responses are shown in circle plot. Red (upregulated) and blue (downregulated) dots in outer circle show the log2(fold change) of genes in each GO term. The color of bar plot in the inner circle is based on z-score and the height of each bar represents the significance of each GO term. p, Canonical pathway perturbations were derived using iPA. Upregulated (red) and downregulated (blue) pathways were identified. GSEA shows inhibition of ‘antigen processing and presentation’ (q) and ‘chemokine signaling pathway’ (r) in the PD-1 Fc treated group (*P < 0.05, **P < 0.01, ****P < 0.0001). Ctl, control; FiTC, fluorescein isothiocyanate.




in vitro with Ova257-264 peptide and selectively cocultured the effec-tor cells with polyclonal Pdl1+/+ or Pdl1−/− CD4+ T cells. Pdl1+/+ T cells were inhibitory, whereas Pdl1−/− T cells were permissive of

cytotoxic T cell (TC) activation (Supplementary Fig. 7a–c). Similarly, polyclonal Pdl1+/+ T cells, but not Pdl1−/− T cells, suppressed Ova-restricted CD4+ OT-II T cell proliferation in response to Ova323-339

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peptide presentation (Supplementary Fig. 7d). These data indicate that Pdl1+/+ T cells are inhibitory to neighboring T cells via the con-ventional PD-L1–PD-1 axis.

To determine the implications of T cell suppression of neighbor-ing effector T cells via the PD-L1–PD-1 axis in cancer, we transferred polyclonal Pdl1+/+ or Pdl1−/− T cells to PDA-bearing Pdl1−/− hosts. Pdl1+/+ T cell transfer resulted in larger tumors compared to transfer of Pdl1−/− T cells and induced intra-tumoral CD4+ and CD8+ T cell suppression (Fig. 6a–c). Analysis of the intra-tumoral immune infil-trate by scRNA-seq indicated a contraction of host CD4+ and CD8+ T cells after Pdl1+/+ T cell transfer compared to Pdl1−/− transfer (Fig. 6d,e). Furthermore, host T cells in tumors of Pdl1+/+ T cell-treated mice expressed lower levels of immunogenic transcription factors (Fos, Nr4a1 and Nr4a2), surface activation markers (Tnfaip3 and Ctla4), inflammatory mediators (Cxcl2 and Il1b), and indicators of cytotoxicity (Gzmb and Klrc1) (Fig. 6f). By contrast, genes that encode calcium binding proteins were downregulated after Pdl1+/+ T cell transfer (S100a6, S100a8 and S100a9). Deeper analysis of transcriptomic data showed that T cells of recipient mice were less enriched for immunogenic upstream regulators—including CD3, p38 MAP Kinase, TNF, and PI3K—after transfer of Pdl1+/+ T cells compared to Pdl1−/− cell transfer (Fig. 6g–i). Collectively, these data imply that Pdl1+/+ T cells induce effector T cell suppression in can-cer even in the absence of host PD-L1. Similarly, adoptive transfer of PDA-infiltrating Pdl1+/+ T cells to Pdl1−/− hosts that were challenged with Ova-expressing PDA resulted in diminished TC expansion and reduced intra-tumoral T cell activation (Fig. 6j).

PD-L1+ T cells induce tolerogenic macrophage differentiation via PD-1 ligation. Ligation of PD-1 in macrophages has recently been shown to decrease their capacity for phagocytosis21. We con-firmed that mouse and human macrophages upregulate PD-1 in PDA (Supplementary Fig. 8a–d), as well as after TLR4 ligation (Supplementary Fig. 8e,f). As we observed that conditional dele-tion of PD-L1 in T cells activated macrophage phenotype in cancer (Fig. 3h–l), we postulated that PD-L1+ T cells interface with mac-rophages via the PD-L1–PD-1 axis and can modulate macrophage differentiation in the TME. Consistent with our hypothesis, we dis-covered that ligation of PD-1 on Pdl1−/− macrophages using PD-L1 Fc promoted an alternatively activated M2-like macrophage pheno-type (Fig. 7a). Treatment of KPC mouse-derived organotypic tumor spheroids (MDOTS) with PD-L1 Fc also induced M2-like polariza-tion of TAMs (Supplementary Fig. 8g). Similar polarizing effects were observed with PD-L1 Fc treatment of human macrophages

(Fig. 7b). By contrast, PD-L1 Fc had no effect on Pd1−/− macro-phages (not shown). To directly test our hypothesis and to determine whether PD-L1-expressing T cells similarly promote tolerogenic macrophage differentiation, we co-incubated polyclonal Pdl1+/+ or Pdl1−/− T cells with Pdl1−/− macrophages. Pdl1+/+ T cells induced a suppressive macrophage phenotype compared to Pdl1−/− T cells (Fig. 7c). Consistent with these observations, TAMs that infiltrated tumors in Pd1−/− mice were markedly more activated than in Pdl1+/+ mice (Supplementary Fig. 8h). Moreover, in vivo adoptive transfer of Pdl1+/+ T cells to PDA-bearing Pdl1−/− mice induced M2-like polarization of TAMs, which was not observed following transfer of Pdl1−/− T cells (Fig. 7d). Analysis of TAMs by scRNA-seq indicated that transfer of Pdl1+/+ T cells reprogrammed the transcriptome of intra-tumoral macrophages and resulted in a less immunogenic TAM phenotype (Supplementary Fig. 8i,j). Consistent with these observations, ingenuity analysis demonstrated that transfer of Pdl1+/+ T cells downregulated M1-associated macrophage signal-ing pathways including IL-6, TNFR, iNOS, HMGB1 and chemo-kine signaling (Fig. 7e). Furthermore, TAMs exhibited diminished enrichment of immunogenic upstream regulators, including IL1β, TNF, IFNγ, TLR4, IL12 and CD38, after Pdl1+/+ T cell adoptive transfer. By contrast, STAT6, FAS, IL10 and SOCS3 were enriched in such cells (Supplementary Fig. 8k,l). Collectively, these data indi-cate that PD-L1-expressing T cells polarize macrophages toward a tolerogenic phenotype in cancer. Notably, the tolerogenic effects of PD-1 ligation on macrophages by PD-L1 Fc was independent of CD80 (Supplementary Fig. 9a,b).

We investigated the signaling mechanisms that regulate PD-1 mediated macrophage polarization. STAT6 signaling, which is associated with M2-like differentiation22, was upregulated in mac-rophages that were cocultured with Pdl1+/+ T cells compared to Pdl1−/− T cells (Fig. 7f). Similarly, Akt signaling, which is also linked to M2-like differentiation23, was upregulated in macrophages after Pdl1+/+ T cell co-culture, whereas p38 MAP Kinase activation was reduced. Treatment of macrophages with PD-L1 Fc induced simi-lar signaling changes (Supplementary Fig. 9c). Immunoblotting experiments confirmed downregulation of M1-associated mark-ers (iNOS, SOCS3) in macrophages and upregulation of Arg1 and pSTAT6 after Pdl1+/+ T cell co-culture (Fig. 7g). Moreover, PD-L1+ T cell effects on macrophage phenotype were STAT6 dependent as Stat6−/− macrophages did not undergo M2-like polarization upon PD-1 ligation (Supplementary Fig. 9d). Further, consistent with their tolerogenic surface phenotype, macrophages entrained by co-culture with Pdl1+/+ T cells were deficient at antigen presentation to

Fig. 5 | PD-L1 signaling in T cells induces STAT3-dependent TH17 differentiation. a,b, Polyclonal Pd1−/− splenic CD4+ T cells were stimulated for 72 h by CD3/CD28 coligation in the presence of PD-1 Fc or igG Fc. CD4+ T cell differentiation was determined by expression of Tbet, GATA3 and Rorγt (a) and iFNγ, iL-17F and iL-17A (b). This experiment was repeated more than five times with similar results. c,d, Polyclonal Pd1−/− CD4+ T cells were stimulated for 72 h by CD3/CD28 coligation in the presence of PD-1 Fc or igG Fc. CD4+ T cell differentiation was determined by qPCR for Ifng, Tnf, Tgfb, Tbet and Gata3 (c), and Rorgt, Il17f and Il17a (d). This experiment was performed in duplicate and repeated twice. e–g Antigen-restricted CD4+ OT-ii T cells were stimulated for 72 h with Ova323–339 peptide in the presence of PD-1 Fc or control igG Fc. T cell differentiation was determined by expression of Tbet (e), Rorγt (f) and FoxP3 (g). This experiment was repeated three times with similar results. h, Polyclonal human PBMC CD4+ T cells were stimulated by CD3/CD28 coligation in the presence of PD-1 Fc or igG Fc for 72 h. Expression of iFNγ, Tbet, iL-17F and iL-17A was determined by flow cytometry. This experiment was performed twice. i, Polyclonal human PBMC CD4+ T cells were stimulated by CD3/CD28 coligation in the presence of PD-1 Fc or igG Fc for 72 h. Expression of Tnf, IL17f, Il17a, LfaFA1, Tgfb and Tbet was determined by qPCR. This experiment was performed in duplicate and was repeated twice. j, Polyclonal Pd1−/− splenic CD4+ T cells were stimulated by CD3/CD28 coligation in the presence of PD-1 Fc or igG Fc. CD4+ T cell expression of pSTAT3, pSTAT1, pAkt, piTK, and pERK was determined by flow cytometry. This experiment was repeated three times with similar results. k, Polyclonal Pd1−/− splenic CD4+ T cells were stimulated by CD3/CD28 coligation in the presence of PD-1 Fc or igG Fc. CD4+ T cell expression of p38, p-p38, Akt, pAkt, SOCS3, FOXO1 and pFOXO1 was determined by immunoblotting. l–n, Polyclonal human PBMC-derived T cells were stimulated by CD3/CD28 coligation in the presence of PD-1 Fc or igG Fc. Cells were harvested at 72 h for analysis by RNA-seq in triplicate. Bubble plots indicate upregulation of STAT3 signaling (l) and inhibition of STAT1 signaling (m) after PD-1 Fc treatment based on iPA of upstream regulators. n, GSEA shows enrichment of FoxO signaling pathway in the PD-1 Fc treated group. o,p, Polyclonal Pd1−/− splenic CD4+ T cells were stimulated by CD3/CD28 coligation in the presence of PD-1 Fc or igG Fc. A small molecule STAT3 inhibitor or vehicle was selectively added to wells. T cell expression of Rorγt (o) and iL-17F (p) was measured at 72 h. This experiment was repeated twice. q, Polyclonal splenic Lck-cre; Stat3fl/fl CD4+ T cells were stimulated by CD3/CD28 coligation in the presence of PD-1 Fc or igG Fc. T cell expression of Rorγt and iL-17F was measured at 72 h (*P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001).




CD4+ T cells compared to macrophages entrained by Pdl1−/− T cells (Fig. 7h–j). Collectively, these data indicate that the PD-L1–PD-1 axis has STAT6-dependent suppressive effects on the macrophage phenotype, which results in impaired adaptive immunity.

DiscussionWe demonstrate that the tolerogenic effects of PD-L1+ T cells are threefold and involve suppressive back-signaling that rivals the

inhibitory effects of PD-1 signaling, suppression of neighboring T cells that is sufficient to promote accelerated tumor growth and intra-tumoral immune tolerance even in the absence of PD-L1 expression from myeloid cells, and induction of M2-like reprogram-ming of TAMs, which also promotes tumor growth and adaptive immune anergy. The PD-1 signaling cascade in T cells and its con-sequences on T cell differentiation have been well-documented. For example, PD-1 signaling promotes Treg differentiation24. However,

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Fig. 6 | PD-L1+ T cells suppress effector T cells in cancer. a–c Pdl1−/− mice were administered orthotopic KPC tumor cells admixed with polyclonal αCD3/αCD28-treated Pdl1+/+ or Pdl1−/− T cells. Tumors were harvested on day 21. a, Representative pictures of PDA tumors (scale bar, 1 cm) and quantitative tumor weight data are shown from four pooled experiments. PDA-infiltrating CD4+ (b) and CD8+ (c) T cells were analyzed by flow cytometry for expression of CD44 and LFA-1. Contour plots and quantitative data from a representative experiment are shown. d–i Pdl1−/− mice were administered orthotopic KPC tumor cells admixed with polyclonal αCD3/αCD28-treated Pdl1+/+ or Pdl1−/− T cells as in (a–c). scRNA-seq was performed on FACS-purified CD45+ tumor-infiltrating leukocytes. d, The distribution of cellular clusters was determined using the t-SNE algorithm. Each cluster is identified by a distinct color. e, A t-SNE plot overlay of tumor-infiltrating leukocytes in tumors of mice treated with Pdl1+/+ T cells (blue) versus Pdl1−/− T cells (red) is shown. Percentage cellular abundance in each cluster is depicted in pie charts. f, Violin plots that compare normalized log expression of selected genes in the T cell cluster for both treatment groups are shown. g, Changes in upstream regulators in host T cells of Pdl1+/+ versus Pdl1−/− T cell-treated mice are shown. Bubble plots indicate downregulation of CD3 signaling (h) and p38 MAP kinase signaling (i) after transfer of Pdl1+/+ T cells compared to Pdl1−/− based on analysis of upstream regulators. j, Pdl1−/− mice were administered orthotopic Ova-expressing KPC tumor cells admixed with PDA-infiltrating Pdl1+/+ or Pdl1−/− T cells and were killed on day 21. Tumor-infiltrating CD8+ T cells were analyzed for the frequency of Ova-pentamer+ cells and expression of CD44, LFA-1 and TNF. Tumor experiments were repeated three times with similar results (*P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001). DC, dendritic cells. Avg. log(FC), average log(fold change).




PD-L1 back-signaling in T cells has not been widely appreciated. Similarly, macrophage programming via ligation of T cell PD-L1 has not been demonstrated. These observations may explain the greater efficacy of PD-L1 blockade compared to PD-1 blockade in select experimental contexts, including that of PDA25,26. In parallel to our finding of PD-L1 back-signaling in T cells, a recent report dem-onstrated PD-L1 signaling in melanoma cells through conserved sequence motifs on the intracellular portion of the PD-L1 cyto-plasmic tail, a process that functionally serves to overcome IFNγ-mediated cytotoxicity27. The oncological implications of PD-L1 signaling in T cells are crystalized by our observation of accelerated spheroid growth and effector T cell suppression after PD-L1 ligation in our PDOTS model of human cancer.

Notably, conditional deletion of PD-L1 in T cells was sufficient to induce adaptive tumor immunity against PDA. We showed that multiple signaling pathways that are linked to T cell immunoge-nicity, including STAT1, AKT, p38 and ERK, are suppressed as a consequence of PD-L1 ligation. Furthermore, we discovered that engagement of PD-L1 promotes TH17 differentiation in a STAT3-dependent manner, including upregulation of Rorγt and IL-17F. In accordance with these results, we found that the osteoarthritis pathway, which is closely linked to TH17 cell expansion, was the most highly upregulated canonical pathway after PD-L1 engage-ment in T cells. However, IL-17A expression was reduced after PD-L1 ligation. Our results therefore suggest that PD-L1 ligation in T cells differentially regulates IL-17A and IL-17F expression. Previous reports have corroborated that IL17A expression is dis-tinctly regulated from IL17F. For example, IL17A expression, but not that of IL17F, is sensitive to inhibition of the PI3K–mTOR path-way28. Consistent with these findings, we found decreased PI3K–mTOR signaling in T cells as a consequence of PD-L1 ligation. Expression of IL-17A and IL-17F by T cells is also differentially regulated in an ITK- and NFATc1-dependent manner. Specifically, CD4+ T cells that are deficient in ITK, a member of the TEC-kinase family, exhibit decreased IL-17A expression, but maintain normal levels of Rorγt and IL-17F expression29. Consistent with these data, we discovered diminished ITK phosphorylation as well as decreased NFAT and TEC-kinase signaling on canonical pathway analysis of T cells treated with PD-1 Fc compared to analysis of non-treated T cells. Notably, whereas IL-17 can have tumor-permissive or tumor-protective properties depending on context30–32, IL-17 sig-naling has been reported to have tumor-promoting effects in PDA, as transformed pancreatic epithelial cells express high IL-17R and proliferate upon its ligation30. Similarly, STAT3 signaling, which we show is required for the PD-L1-mediated TH17 phenotype, has been linked to accelerated tumorigenesis in PDA33,34. Thus, in addition to mitigating cellular activation in T cells, engagement of

PD-L1 can potentially promote oncogenic progression as a result of the collateral consequences of upregulated IL-17F expression. It is noteworthy that we discovered that, compared to PD-L1− CD4+ T cells, a higher percentage of PD-L1+ cells express IL-17, as well as transcription factors and cytokines that are indicative of TH1 and TH2 cells. The latter observation is particularly interesting consid-ering that engagement of PD-L1 skews CD4+ T cells toward a TH17-like phenotype. However, as these experiments were performed with unstimulated splenic T cells, these differences may just reflect their baseline activation state.

In addition to the discovery of fate-determining PD-L1 back-signaling in T cells, our parallel observation that PD-L1-expressing T cells suppress neighboring PD-1+ T cells has critical implications for immune-suppression in the TME. Indeed, the transfer of poly-clonal Pdl1+/+ T cells to Pdl1−/− hosts that were challenged with PDA was sufficient to induce larger tumors than transfer of Pdl1–/– T cells and marked host T cell suppression. Pdl1+/+ T cells corrupted the T cell antigen receptor, p38 MAP kinase, and PI3K–Akt signaling pathways in effector T cells. These data indicate that intra-tumoral conventional T cells potently suppress neighboring T cells via the PD-L1–PD-1 axis in cancer. In a similar study, we recently reported that high levels of checkpoint ligand expression on γδ T cells, including PD-L1 and galectin-9, powerfully inhibit the activation of adjacent αβ T cells in PDA via ligation of their respective check-point receptors25. Furthermore, we previously demonstrated greater spatial proximity between PD-L1+ T cells and PD-1+ T cells as compared to expanded distances between PD-L1+ APC and PD-1+ T cells in the PDA TME25. Consequently, T cells may be more effi-cient suppressors of effector T cells via the PD-L1–PD-1 axis in situ compared to APC.

Despite the fact that T cells and TAMs have less opportunity for direct cellular interface, we have previously demonstrated that the character and volume of infiltrating T cells in PDA is to a large extent dictated by the phenotype of TAMs8,35. Therefore, TAMs are an attractive potential target in PDA immunotherapy. M1-like TAMs promote immunogenic T cell differentiation, whereas M2-like TAMs generate TH2 cells and Tregs (ref. 36). However, the drivers of macrophage polarization in the PDA TME are incom-pletely understood. We demonstrate in tumor models that ligation of PD-1 in macrophages by PD-L1 promotes the differentiation of tolerogenic macrophages, which in turn both suppresses T cell acti-vation and promotes tumor growth. Furthermore, Pdl1+/+ T cells induced TAMs to exhibit diminished enrichment of immunogenic regulators, including IL1β, TNF, IFN-γ and IL12. By contrast, IL-10, FAS and STAT6 were enriched in TAMs after PD-1 ligation. The tolerogenic effects of PD-1 signaling in macrophages were STAT6-dependent. More broadly, these observations imply that signaling

Fig. 7 | PD-L1+ T cells promote suppressive macrophage differentiation. a, Lipopolysaccharide (LPS)-activated Pdl1−/− splenic macrophages were treated with PD-L1 Fc or igG Fc and tested for expression of cytokines and activation markers at 24 h by qPCR. This experiment was performed in duplicate and was repeated three times with similar results. b, LPS-activated PBMC-derived human macrophages were treated with PD-L1 Fc or igG Fc and tested for expression of cytokines and activation markers at 24 h by flow cytometry. This experiment was repeated twice. c, Polyclonal splenic Pdl1+/+ or Pdl1−/− T cells were prestimulated for 24 h by CD3/CD28 ligation and then cocultured with LPS-activated Pdl1−/− macrophages. Macrophage phenotype was analyzed at 24 h. This experiment was repeated three times with similar results. d, Pdl1−/− mice were administered orthotopic KPC tumors admixed with Pdl1+/+ or Pdl1−/− T cells (n = 5 per group). Tumors were harvested on day 21. Schematic of experimental design is shown. TAMs were tested for expression of CD38 and MHC ii. This experiment was repeated twice with similar results. e, Pdl1−/− mice were administered orthotopic KPC tumors admixed with Pdl1+/+ or Pdl1−/− T cells. Tumors were harvested on day 21 and scRNA-seq was performed on FACS-purified CD45+ tumor-infiltrating leukocytes. iPA-based canonical pathway perturbations in the CD68hi macrophage cluster for the Pdl1+/+ versus Pdl1−/− T cell transfer groups were identified. Upregulated (red) and downregulated (blue) pathways are shown. f, Splenic Pdl1+/+ or Pdl1−/− T cells were prestimulated by CD3/CD28 ligation and then cocultured with Pdl1−/− macrophages. Macrophage expression of pSTAT3, pSTAT6, pAKT and p-p38 MAPK was determined at 24 h. This experiment was repeated three times with similar results. g, Pdl1−/− macrophages were treated with PD-L1 Fc or igG Fc. Macrophage expression of Arg1, iNOS, SOCS3, STAT6 and pSTAT6 was determined by immunoblotting. Loading controls are shown for each membrane. h–j, Pdl1−/− splenic macrophages were cocultured with PDA-infiltrating Pdl1+/+ and Pdl1−/− T cells for 24 h. Macrophages were then purified, loaded with Ova323–339 peptide, and used to stimulate Ova-restricted CD4+ T cells. T cell expression of CD6 (h), CD44 (i) and Tbet (j) was determined at 72 h. This experiment was repeated three times with similar results (*P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001).




in the immunological synapse between APC and T cells is bidirec-tional, and that T cells provide critical fate-determining signals to APC. Our findings of significant PD-1 signaling in macrophages are consistent with a recent report, which showed that PD-1 ligation in macrophages inhibits their capacity for phagocytosis21. In contrast to the suppressive effects of PD-L1+ T cells on TAMs, previous work has shown that PD-L1 expression on T cells is necessary for DC maturation in the context of influenza infection, which suggests that effects may be contingent on both the environmental context and

the APC subtype37. Notably, we found that macrophage expression of PD-1 is upregulated by ligation of TLR4. We previously demon-strated that PDA tumors are rife with both sterile and microbial-derived TLR4 ligands38–40. Furthermore, the PDA microbiome, and specifically TLR4 ligation, induces crippling M2-like macrophage differentiation in situ39. Our current work may provide a mechanis-tic link to the latter observation, as microbial-derived TLR4 signals upregulate PD-1 in macrophages, which induces an M2-like pro-gram upon activation by PD-L1. However, the causal link between

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the PDA microbiome, PD-1 signaling and suppressive macrophage polarization requires further detailed study. In summary, our work demonstrates that T cell expression of PD-L1 has pleiotropic effects on innate and adaptive immune tolerance in cancer. These findings elucidate our understanding of cellular crosstalk at the immunolog-ical synapse and signaling in the TME and may have critical impli-cations for immunotherapeutic response and resistance in patients with cancer.

online contentAny methods, additional references, Nature Research reporting summaries, source data, extended data, supplementary informa-tion, acknowledgements, peer review information; details of author contributions and competing interests; and statements of data and code availability are available at https://doi.org/10.1038/s41590-020-0620-x.

Received: 16 January 2019; Accepted: 24 January 2020; Published online: 9 March 2020

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MethodsAnimals and in vivo procedures. C57Bl/6J (H-2Kb), Ifng−/−, Il27r−/−, Lck-cre, Cd4-cre, Stat3fl/fl, Pd1−/−, Stat6−/−, OT-I, and OT-II mice were purchased from The Jackson Laboratory. Pdl1−/− mice were a gift of C. Drake (Columbia University, NY). Pdl1fl/fl mice were a gift of T. Wang (Columbia University, NY). KC mice were a gift of D. Bar-Sagi (New York University, NY). Animals were housed in a clean vivarium and were fed standard mouse chow. Both male and female mice were used, but animals were gender-matched within each experiment. For orthotopic pancreatic tumor challenge, 8–10-week-old mice were administered intra-pancreatic injections of FC1242 tumor cells derived from KPC mice, as described previously38. In selected experiments we used FC1242 cells that we engineered to express ovalbumin using the pCIneo-OVA vector (Addgene) as we have described8. PDA cells were suspended in PBS with 50% Matrigel (BD Biosciences) and 1 × 105 tumor cells were injected into the body of the pancreas by laparotomy. Mice were killed three weeks later for analysis. In selected experiments T cells (1:5 ratio) or macrophages (1:4 ratio) were admixed with tumor cells before orthotopic administration. In other experiments, tumor bearing mice were treated with TLR2 ligand (Pam3CSK4, 50 μg, i.p., 3x per week; Invivogen). In our liver metastases model, KPC-derived tumor cells were suspended in PBS and 1 × 106 tumor cells were injected into the portal venous system, as we have described8. In selected experiments, mice were challenged with B16 melanoma (gift of R. DeMatteo, University of Pennsylvania) or MCA38 colon cancer (both 1 × 106 cells, SQ; gift of A. Frey, New York University). Chronic pancreatitis was induced using a regimen of seven-hourly injections of caerulein (50 μg kg−1, i.p.; Sigma) thrice weekly for three weeks, as we have described41. Animal procedures were approved by the New York University School of Medicine Institutional Animal Care and Use Committee.

Murine and human cellular isolation, flow cytometry and FACS. Single-cell suspensions of mouse PDA tumors were prepared for flow cytometry as described previously35. In brief, pancreata were placed in cold 2% FACS (PBS with 2% FBS) with Collagenase IV (1 mg ml−1; Worthington Biochemical), trypsin inhibitor (1 mg ml−1; EMD Millipore), and DNase I (2 U ml−1; Promega), and were minced with scissors to pieces less than 1mm in size. Tissues were then incubated at 37 °C for 20 min with gentle shaking every 5 min. Specimens were passed through a 70 μm mesh and centrifuged at 350 g for 5 min. Splenocytes were prepared by manual disruption and splenic T cells or macrophages were isolated by positive selection using magnetic beads (Miltenyi) as described previously25. BMDM were prepared and cultured as described previously42. Cell pellets were resuspended and cell labeling was performed after blocking FcγRIII/II with a mAb to CD16/CD32 (eBioscience) by incubating 1 × 106 cells with 1 μg of fluorescently conjugated mAbs directed against mouse, PD-L1 (10F.9G2), CD3 (17A), CD4 (RM4-5), CD8 (53-6.7), CD45 (30-F11), 4-1BB (17B5), LAG-3 (C9B7W), CD44 (IM7), CD206 (C068C2), CD39 (Duha5), CD49b (DX5), CD73 (TY/11.8), CD11b (M1/70), CD11c (N418), MHC II (M5/114.15.2), IL-4 (11B11), IL-10 (JES5-16E3), IL-17F (8F5.1A9), IL-17A (TC11-18H10.1), IFN-γ (XMG1.2), TNF (MP6-XT22), ICOS (15F9), CD69 (H1.2F3), CD107a (1D4B), PD-1 (29F.1A12), CD25 (3C7), LFA-1 (H155-78), OX40 (OX-86), CD38 (90), CD80 (16-10A1), GATA3 (16E10A23), Tbet (4B10), FOXP3 (MF-14), EOMES (7C9B03), pERK (6B8B69), pSTAT1 (10C4B40), pSTAT3 (13A3-1), pSTAT4 (15A1B41), pSTAT6 (A15137E), TGF-β (TW7-16B4), Ms IgG (Poly4053), Rt IgG (Poly4054; all BioLegend), iNOS (CXNFT; Thermo Fisher Scientific), Rorγt (Q31-378; BD Biosciences), pAKT (545007; R&D Systems), pSTAT5 (47/Stat5[pY694]; BD Biosciences), pBTK/ITK (M4G3LN; Thermo Fisher Scientific) and p38MAPK (36/p38; BD Biosciences). Ova-pentamer staining was performed using Pro5 MHC Class I pentamer (ProImmune). For phosphoflow staining, cells were fixed with 1.85% formaldehyde in PBS for 7 min at 37 °C, followed by True-Phos Perm Buffer (Biolegend) as per the manufacturer’s protocol. Human pancreatic tumor and PBMC were collected under an institutional review board (IRB)-approved protocol. Human pancreatic leukocytes were prepared in a similar manner to mice. PBMC were isolated by overlaying whole blood diluted 1:1 in PBS over an equal amount of Ficoll (GE Healthcare, Princeton, NJ). Cells were then spun at 2100 RPM and the buffy coat harvested as we have described43. Analysis of human cells was performed using fluorescently conjugated antibodies directed against CD45 (HI30), CD3 (SK7), CD62L (DREG-56), CD44 (BJ18), CD69 (FN50), ICOS (C398.4A), TNF (Mab11), LFA-1 (m24), Granzyme B (QA16A02), Tbet (4B10), IL-17F (Poly5166), CD68 (Y1/82A), CD80 (2D10), CD86 (BU63), IFN-γ (4S.B3), CD163 (GHI/61, all BioLegend). Dead cells were excluded from analysis using zombie yellow (BioLegend). Flow cytometry was performed on the Attune NxT Acoustic Focusing Cytometer (Thermo Fisher Scientific). FACS-sorting was performed on the SY3200 (Sony). Data were analyzed using FlowJo (Tree Star).

T cell activation assays and macrophage polarization experiments. For antibody-based T cell proliferation assays, mouse splenic or human PBMC-derived T cells were activated using CD3/CD28 coligation in 96 well plates, as we described previously25. In selected experiments neutralizing mAbs specific to IL-6Ra (D7715A7; BioLegend,), IL4 (11B11; BioLegend), IFNγ (XMG1.2; BioLegend), IL27 p28 (MM27-7B1; eBioscience), IL17A (TC11-18H10.1; BioLegend), IL10 (JES5-2A5; BioLegend), or small molecule inhibitors against STAT3 (cucurbitacin I, 0.5 uM; Tocris), mTOR (rapamycin, 5 nM; Tocris), Akt (AZD5364, 5 nM;

Selleck Chemicals) or PI3K (busparlisib, 5 nM; Selleck Chemicals) were used. For antigen-restricted CD4+ T cell stimulation assays, splenic OT-II T cells were cultured directly with Ova323-339 peptide (10 µg ml−1) or with macrophages pulsed with Ova323-339 peptide in a 5:1 ratio. For CD8+ T cell activation assays, splenic OT-I T cells were cultured with Ova257-264 peptide (10 µg ml−1). T cell activation was determined at 72 h by flow cytometry. In selected wells, T cells were treated with PD-1 Fc or control IgG Fc (2.5 µg ml−1; R&D Systems). In other experiments, T cells were treated with an αPD-L1 mAb (10 µg ml−1, 10F.9G2) or Rat IgG2bκ (both BioLegend). Alternatively, in some experiments polyclonal T cells were added to OT-I or OT-II stimulation assays in a 1:5 ratio. For mouse or human macrophage polarization assays, T cells were cocultured with naïve splenic or PMBC-derived macrophages for 36 h in a 1:4 ratio. PD-L1 Fc (2.5 µg ml−1; R&D Systems) was selectively used to treat macrophages. In some experiments, a neutralizing αCD80 mAb was used (0.5 µg ml−1; R&D).

Immunoblotting. For protein extraction, tissues were homogenized in ice-cold RIPA buffer. Total protein was quantified using the DC Protein Assay according to the manufacturer’s instructions (BioRad). Immunoblotting was performed as described previously, with minor modifications38. Briefly, 10% Bis-Tris polyacrylamide gels (NuPage, Invitrogen) were equiloaded with 10–30 μg of protein, electrophoresed at 200 V, and electrotransferred to PVDF membranes. After blocking with 5% BSA, membranes were probed with primary antibodies to Arg1 (AF5868; R&D Systems), SOCS3 (ab16030), PD-L1 (ab238697; both Abcam), p38MAPK (#9212), p-p38MAPK (#9216), AKT (#9272), pAKT (#4060), FOXO1 (#2880), pFOXO1 (#9461), iNOS (#13120), STAT6 (#9362), pSTAT6 (#56554), vinculin (#13901), and β-actin (#3700; all Cell Signaling). Blots were developed by ECL (Thermo Fisher Scientific).

qPCR. For qPCR, total RNA was extracted using an RNeasy mini kit (Qiagen) and cDNA was synthesized using the iScript cDNA Synthesis Kit (BioRad). Real-time qPCR was performed using the BioRad Real-Time PCR System. Each reaction mixture contained 10 μl of SsoAdvanced Universal SYBR Green Supermix (BioRad), 0.5 μl of forward and reverse primers (Invitrogen), and 3 μl of cDNA (corresponding to 50 ng of RNA). The qPCR conditions were: 50 °C for 2 min, 95 °C for 10 min, followed by 40 cycles at 95 °C for 15 s, and 60 °C for 1 min. Amplification of specific transcripts was confirmed by melting curve profiles generated at the end of the PCR program. Expression levels of target genes were normalized to the expression of the 18S gene (as an internal control) and were calculated based on the comparative cycle threshold method (2-ΔΔCt). The mouse primer sequences used in the study were as follows: Tgfb: F- TGACGTCACTGGAGTTGTACGG, R- GGTTCATGTCATGGATGGTGC; Cd38: F- TTGCAAGGGTTCTTGGAAAC, R- CGCTGCCTCATCTACACTCA; Cd64: F- ATTCGGAGGTCGCCATTCTGA, R- CCATCGCTTCTAACTTGCTGA; Cd68: F- ACTTCGGGCCATGTTTCTCT, R- GCTGGTAGGTTGATTGTCGT; Inos: F- AGCCTAGTCAACTGCAAGAG, R- TCTTGTATTGTTGGGCTGAGA; Tnf: F- CTACTGAACTTCGGGGTGAT, R- CTTGGTGGTTTGTGAGTGTG; Il10: F- CACAAAGCAGCCTTGCAGAA, R- AGAGCAGGCAGCATAGCAGTG; Ifng: F- CGGCACAGTCATTGAAAGCCTA, R- GTTGCTGATGGCCTGATTGTC; Cd86: F- TCCAGAACTTACGGAAGCACCCACG, R- CAGGTTCACTGAAGTTGGCGATCAC; Tbet: F- GCCAGGGAACCGCTTATATG, R- GACGATCATCTGGGTCACATTGT; Rorgt: F- GCAGCGCTCCAACATCTTCT, R- ACGTACTGAATGGCCTCGGT; Il17f: F- ACGTGAATTCCAGAACCGCT, R- TTGGAGATCGGGCTTCACAC; Il17a: F-GCTCCAGAAGGCCCTCAGA R-CTTTCCCTCCGCATTGACA; Gata3: F- CCAAGGCACGATCCAGCACAGA, R- TGCCGACAGCCTTCGCTTGG; H2ab: F- AGCCCCATCACTGTGGAGT, R- GATGCCGCTCAACATCTTGC.

The human primer sequences used in the study were as follows: TNF: F- ATGAGCACTGAAAGCATGATCC, R- GAGGGCTGATTAGAGAGAGGTC; IL17F: F- TGCCAGGAGGTAGTATGAAGCTT, R- ATGCAGCCCAAGTTCCTACACT; IL17A: F- TCCCACGAAATCCAGGATGC, R- GGATGTTCAGGTTGACCATCAC; LFA1: F- AAATGGAAGGACCCTGATGCTC, R- TGTAGCGGATGATGTCTTTGGC; TGFB: F- CACCCGCGTGCTAATGG, R- ATGCTGTGTGTACTCTGCTTGAACT; TBET: F- GCCTACCAGAATGCCGAGATTA, R- GGACTCAAAGTTCTCCCGGAAT; GATA3: F- ACCACAACCACACTCTGGAGGA, R- TCGGTTTCTGGTCTGGATGCCT.

Sequential immunohistochemistry and image acquisition. For histological analysis, human PDA tissues were fixed with 10% buffered formalin, dehydrated in ethanol, and embedded with paraffin and sectioned at a thickness of 5 um. Multiplex immunofluorescence was performed on a Leica Bond Rx automated stainer, according to established manufacturer protocols for sequential antibody binding and amplification (Leica). Primary antibodies against CD3 (LN10, undiluted; Leica), CD163 (MRQ-26, 1:100; Roche), Pan-cytokeratin (AE1 and AE3, 1:40; Agilent) and either PD1 (D4W2J, 1:100; Cell Signaling) or PD-L1 (22C3, 1:40; Agilent) were incubated sequentially, followed by a cocktail of horse radish peroxidase-conjugated secondary antibodies against mouse and rabbit immunoglobulins (PerkinElmer). After tyramide-mediated signal amplification

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and covalent linkage of the individual Opal fluorophores to the relevant epitope or epitopes, the primary and secondary antibodies were removed and the next cycle of immunostaining was initiated. The sequence of Panel 1 was PD1/Opal 570, CD3/Opal 650, CD68/Opal 520, and PanKeratin/Opal 690. The Panel 2 sequence was the same, except PD-L1 replaced PD1. All slides were counterstained with spectral DAPI (FP1490; PerkinElmer). Monoplex controls were used to confirm appropriate staining for all antibodies integrated into the multiplex panels. Multispectral imaging was performed on a Vectra 3 system (PerkinElmer). Slides were imaged at ×200 magnification and the fluorophore signals were captured by a multispectral camera module. The fluorophore emission signatures were spectrally unmixed and the images were analyzed with InForm software (PerkinElmer). Autofluorescence, obtained from an unstained slide, was removed from the composite images and pseudo-colored images were exported as TIF files.

Conventional RNA-seq and analysis. RNA-seq libraries were prepared using the Illumina TruSeq Stranded Total RNA library prep, after ribodepletion with Ribozero Gold kit (cat no. 20020597; Illumina) starting from 500 ng of DNAse I-treated total RNA, following the manufacturer’s protocol, with the exception that 9 cycles of PCR were performed to amplify the libraries. The amplified libraries were purified using AMPure beads, quantified by Qubit and qPCR, and visualized in an Agilent Bioanalyzer (Agilent). The libraries were pooled equimolarly, and were sequenced on one lane of an Illumina HiSeq 2500 flow cell. The raw FASTQ reads were aligned to the mm10 mouse reference genome using STAR aligner44. FastQ Screen was used to check for any contamination in the samples and Picard RnaSeqMetrics was used to obtain the metrics of all aligned RNA-seq reads. The featureCounts program was used to quantify the gene expression levels45. The raw gene counts data was used for further differential expression analysis. To identify the differentially expressed genes, the DESeq2 R package was used46. The resulting genes with adjusted P < 0.05 were considered significant. Heatmaps were generated using the pheatmap R package. To determine the functional annotation of the genes that were expressed at a significant level, GO analysis was performed and the expression levels of genes in each term were represented as a GO circle plot using the R package GOplot. To identify the signaling pathways in which the genes are enriched, IPA was carried out for genes whose expression was considered to be significant. The regulatory network of genes that are associated with signaling pathways was represented using Cytoscape47. GSEA was performed on differentially expressed genes. Upregulated and downregulated sets of genes were ranked on the basis of their average and normalized log2(fold change) between the treatment and control group, and each gene set was assessed for enrichment in the Kyoto Encyclopedia of Genes and Genomes (KEGG)_2016 geneset library [http://amp.pharm.mssm.edu/Enrichr/#stats] using the Python package gseapy [https://pypi.org/project/gseapy] for the analyses.

scRNA-seq. Sequencing results were demultiplexed and converted to FASTQ format using Illumina bcl2fastq software. The Cell Ranger single-cell software suite (https://support.10xgenomics.com/single-cell-gene-expression/software/pipelines/latest/what-is-cell-ranger) was used to perform sample demultiplexing, barcode processing, and single-cell 3′ gene counting. The cDNA insert was aligned to the mm10 reference genome, assembly GRCm38. Only confidently mapped non-PCR duplicates with valid barcodes and unique molecular identifiers were used to generate the gene–barcode matrix. To account for technical batch differences, we utilized the Seurat alignment method for data integration. We took the union of the top 2,000 genes with the highest dispersion from both datasets and ran a canonical correspondence analysis (CCA) to determine the common sources of variation between datasets. We next aligned the subspaces based on the first 16 canonical correlation vectors, to generate a new dimensionality reduction that was then used for further analysis. The data were visualized with t-distributed Stochastic Neighbor Embedding (t-SNE) based on the aligned CCA. Marker genes were determined on the basis of differential expression analysis using a Wilcoxon rank sum test for each cluster. Cell type identities of known population markers were assigned as follows: CD68hi macrophages: Cd68hiCsf1rhiFcgr1hiAdgre1hiItgamhi; T cells: Cd3ghiCd3dhiCd3ehiCd8ahi; CD103hi MHCIIhi DC: ItgaehiH2-Ab1hiIgamlo; B cells: Cd19hiCd79ahiCd79bhiCd3elo; CX3CR1hi macrophages: Cx3cr1hiAdgre1hiArg1hiItgamhi; MDSC: Cd177hiCxcr2hiMmp8hiArg2hiItgamhi; Batf3hi MHCIIhi DC: Batf3hiH2-Ab1hiCd74hi.

MDOTS and PDOTS preparation, treatment and analysis. MDOTS and PDOTS were prepared as described previously, with slight modifications8,48. Briefly, mouse orthotropic PDA tumors or human surgically resected tumor specimens were received fresh in DMEM medium on ice and were minced in 10 cm dishes. Minced tumors were resuspended in DMEM + 10% FBS with 100 U ml−1 collagenase type IV to obtain spheroids. Partially digested samples were pelleted and then resuspended in fresh DMEM + 10% FBS and were strained over both 100 mm and 40 mm filters to generate S1 (>100mm), S2 (40–100mm), and S3 (<40mm) spheroid fractions, which were subsequently maintained in ultra-low-attachment tissue culture plates. An aliquot of the S2 fraction was pelleted and resuspended in type I rat tail collagen and the spheroid–collagen mixture was then injected

into the center gel region of the DAX-1 3D microfluidic cell culture chip (AIM Biotech). After 30 min at 37 °C, collagen hydrogels containing MDOTS or PDOTS were hydrated with media with indicated treatments (see ‘Engagement of PD-L1 by PD-1 induces inhibitory signaling in T cells and drives TH17 differentiation’). Spheroids were harvested on day 3 for analysis by flow cytometry. Images were captured on a Nikon Eclipse 80i fluorescence microscope equipped with Z-stack (Prior) and CoolSNAP CCD camera (Roper Scientific). Image capture and analyses were performed using the NIS-Elements AR software package (Nikon). Human biological samples were sourced ethically, and their research use was in accordance with the terms of informed consent under an IRB-approved protocol.

Lentiviral transfection. Mouse PD-L1 [NM_021893.3] and eGFP cDNAs were cloned into the pLV-Lentivirus gene expression vector obtained from Vector Builder. Mutations were introduced by using the QuikChange II site-directed mutagenesis kit (Agilent). Viruses were produced by cotransfection of HEK293T cells with lentiviral constructs and the packaging vectors psPAX2 (Addgene, cat no. 12260) and pMD2.G (Addgene, cat no. 12259). At 48 h after transfection, the cell culture media were filtered through a 0.45 mm filter, and the viral supernatants were supplemented with polybrene (8 µg ml−1; Sigma) were used for infections. Mouse Pdl1−/− T cells were infected for 48 h, at 37 °C. Subsequently, the cells were harvested and analyzed for PD-L1 expression.

Statistical analysis. Data is presented as mean ± s.e.m. Statistical significance was determined by the Student’s t test using GraphPad Prism 6 (GraphPad). P values < 0.05 were considered to be significant. Significance for the GSEA analysis and the differential gene expression based on scRNA-seq was determined using the Wilcoxon rank sum test with Bonferroni multiple-comparison correction. Comparisons for more than two groups were calculated using two-way ANOVA followed by Bonferroni multiple-comparison correction.

Reporting Summary. Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Data availabilitySequence data are available in the Gene Expression Omnibus (GEO) database at NCBI under accession code GSE145905.

References 41. Bedrosian, A. S. et al. Dendritic cells promote pancreatic viability in mice

with acute pancreatitis. Gastroenterology 141, 1915–1926.e14 (2011). 42. Greco, S. H. et al. Mincle suppresses Toll-like receptor 4 activation. J. Leukoc.

Biol. 100, 185–194 (2016). 43. Rehman, A. et al. Role of fatty-acid synthesis in dendritic cell generation and

function. J. Immunol. 190, 4640–4649 (2013). 44. Dobin, A. et al. STAR: ultrafast universal RNA-seq aligner. Bioinformatics 29,

15–21 (2013). 45. Liao, Y., Smyth, G. K. & Shi, W. featureCounts: an efficient general purpose

program for assigning sequence reads to genomic features. Bioinformatics 30, 923–930 (2014).

46. Love, M. I., Huber, W. & Anders, S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 15, 550 (2014).

47. Shannon, P. et al. Cytoscape: a software environment for integrated models of biomolecular interaction networks. Genome Res. 13, 2498–2504 (2003).

48. Jenkins, R. W. et al. Ex vivo profiling of PD-1 blockade using organotypic tumor spheroids. Cancer Discov. 8, 196–215 (2017).

AcknowledgementsWe acknowledge the use of the Genome Technology, Experimental Pathology and Flow Cytometry core facilities at NYU School of Medicine. These shared resources are partially supported by the Cancer Center Support Grant, P30CA016087, at the Laura and Isaac Perlmutter Cancer Center. The Vectra3 imaging system was purchased through NIH Shared Instrument Grant S10 OD021747. This work was supported by the American College of Surgeons Resident Research Fellowship (B.D.) and NIH grants CA168611 (G.M.), CA203105 (G.M.), CA215471 (G.M.), CA19311 (G.M.) and DK106025 (G.M.).

Author contributionsB.D. and S.A. prepared the manuscript, performed in vivo and in vitro experiments and data analysis, and designed, supervised and interpreted the study; M.F.C., G.S., R.D.S., D.W., C.F., K.H.T. and T.C. performed in vitro experiments; M.L, C.B, E.L., B.S., R.C., J.W. and Z.S. performed in vivo experiments in addition to manuscript and figure preparation; M.K. performed data analyis; B.A., M.S.F., S.N., J.A.K.R., E. Kruger, M.I.U.H. and J.K. performed in vivo experiments, data analysis and manuscript review; W.W., M.H. S.A.A.S, G.B., G.W., K.-K.W., J.L. and E. Kurz performed data analysis and critical review; S.S. and C.L. performed immunofluorescence; G.M. conceived, designed,


http://amp.pharm.mssm.edu/Enrichr/#stats

http://amp.pharm.mssm.edu/Enrichr/#stats

https://pypi.org/project/gseapy

https://pypi.org/project/gseapy

https://support.10xgenomics.com/single-cell-gene-expression/software/pipelines/latest/what-is-cell-ranger

https://support.10xgenomics.com/single-cell-gene-expression/software/pipelines/latest/what-is-cell-ranger

https://www.ncbi.nlm.nih.gov/nuccore/NM_021893.3

https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE145905


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supervised, analyzed and interpreted the study and provided critical review, and is senior author and corresponding author.

Competing interestsG.M. has research agreements with GSK, Pfizer and Puretech Health. K.K.W. is a founder and equity holder of G1 Therapeutics and has sponsored research agreements with MedImmune, Takeda, TargImmune, Bristol-Myers Squibb, Mirati, Merus and Alkermes, and has consulting and sponsored research agreements with AstraZeneca, Janssen, Pfizer, Novartis, Merck, Ono and Array. The remaining authors declare no competing interests.

Additional informationSupplementary information is available for this paper at https://doi.org/10.1038/s41590-020-0620-x.

Correspondence and requests for materials should be addressed to G.M.

Peer review information Zoltan Fehervari was the primary editor on this article and managed its editorial process and peer review in collaboration with the rest of the editorial team.

Reprints and permissions information is available at www.nature.com/reprints.


https://doi.org/10.1038/s41590-020-0620-x

https://doi.org/10.1038/s41590-020-0620-x

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