Abstract
Natural killer (NK) cells are critical mediators of host immunity to pathogens. Here, we demonstrate that the ER stress sensor inositol-requiring enzyme 1 (IRE1α) and its substrate transcription factor X-box-binding protein 1 (XBP1) drive NK cell responses against viral infection and tumors in vivo. IRE1α-XBP1 were essential for expansion of activated mouse and human NK cells and are situated downstream of the mTOR signaling pathway. Transcriptome and chromatin immunoprecipitation analysis revealed c-Myc as a novel and direct downstream target of XBP1 for regulation of NK cell proliferation. Genetic ablation or pharmaceutical blockade of IRE1α downregulated c-Myc, and NK cells with c-Myc haploinsufficency phenocopied IRE1α-XBP1 deficiency. c-Myc overexpression largely rescued the proliferation defect in IRE1α−/− NK cells. Like c-Myc, IRE1α-XBP1 also promotes oxidative phosphorylation in NK cells. Overall, our study identifies a novel IRE1α-XBP1-cMyc axis in NK cell immunity, providing new insight into host protection against infection and cancer.
Introduction
NK cells are critical mediators of host immunity against malignancies and viral infection1. Although extrinsic regulators of NK cell development and function including diverse ligands of key NK cell receptors and proinflammatory cytokines from the microenvironment have been identified2, 3, 4, we understand relatively little about how NK cells translate these signals into critical effector functions5, 6. Furthermore, novel features of NK cells have emerged in recent years, including their ability to undergo clonal proliferation and generate long-lived memory; however, the molecular mechanisms underlying these “adaptive” properties require further characterization. The emerging interest in developing NK cell-based cancer immunotherapy7 and new vaccine strategies for controlling lethal infectious diseases highlights an urgent need for identifying novel intrinsic regulators of NK cell-mediated immunity.
The activation of ER stress sensor IRE1α and its substrate transcription factor XBP18,9 is a hallmark of ‘professional’ secretory cells that must constitutively deal with a high demand for protein synthesis, folding, and secretion10,11,12. This highly evolutionarily conserved signaling pathway is also activated in tumor cells13 and in myeloid-derived suppressor cells14, macrophages15, T cells16 and dendritic cells17 in response to external stimuli such as hypoxia, nutrient-deprivation and low pH. However, it is unknown whether NK cell function is driven by IRE1α/-XBP1 signaling, and if so, what specific activities it controls.
To address these questions, we used an IRE1α activation reporter mouse strain to demonstrate that activated NK cells upregulate the IRE1α-XBP1 pathway following exposure to pro-inflammatory cytokines in vitro and to viral infection in vivo. We use newly-engineered genetic mouse models to identify the IRE1α/XBP1 pathway as a positive cell-intrinsic regulator of NK cell proliferation and expansion during viral infection or lymphopenia, and of NK cell-mediated anti-tumor protection. IRE1α-XBP1 is induced by mTOR and STAT4 signaling pathways in activated NK cells. XBP1 facilitates NK cell expansion in part by directly binding to and activating the c-Myc promoter to upregulate key c-Myc target genes required for NK cell expansion as well as by controlling mitochondrial respiration. This study reveals an unexpected role for the IRE1α/XBP1 pathway and for the transcriptional regulator c-Myc in NK cell-mediated immunity.
Results
Upregulation of the IRE1α/XBP1 pathway in activated NK cells
We tested whether the IRE1α-XBP1 ER Stress Response (or Unfolded Protein Response, UPR) is induced in activated NK cells in vivo following mouse cytomegalovirus (MCMV) exposure, where NK cells are critical for infection control2, 18. Unbiased transcriptome analysis highlighted ER stress as a top enriched gene ontology (GO) category in MCMV-primed NK cells (Fig. 1a), with rapid upregulation at day 1.5 post infection (PI) (Fig. 1b), accompanied by generation of the active spliced XBP1s RNA, and upregulation of canonical XBP1 target genes (Supplementary Fig. 1a). Consistent with this, transgenic ER stress activated indicator (ERAI) reporter mice19 revealed minimal IRE1α activation in naïve NK cells, but significantly elevated levels in activated NK cells at day 2 PI that returned to baseline by day 7 PI (Fig. 1b,c) indicating transient activation of this pathway in response to viral infection.
NK cell activation during viral infection occurs when exposed to proinflammatory cytokines including IL-12 and IL-18, and ligation of activating receptors4, 6, 20, 21. Consistent with in vivo findings, RNA-seq analysis of IL-12 and IL-18 in vitro activated NK cells showed robust upregulation of the IRE1α-XBP1 UPR signature, Xbp1s induction and activation of canonical XBP1 target genes (Fig. 1d, Supplementary Fig. 1b). Results using the ERAI reporter mouse confirmed IRE1α activity in cytokine-activated NK cells from spleen and bone marrow (BM) (Fig. 1e). Notably. XBP1 activation was also observed in primary human NK cells following IL-12 and IL-18 stimulation in vitro (Fig. 1f).
We next identified both Stat4 and the mammalian target of rapamycin (mTOR) as upstream regulators of IRE1α-XBP1 function in infection and in cytokine-activated NK cells. STAT4−/− NK cells displayed reduced Xbp1 and downstream target gene activation during MCMV infection (Supplementary Fig. 1c), and pharmaceutical blockade of mTOR in NK cells significantly reduced IRE1α activation in response to cytokine stimulation in vitro (Supplementary Fig. 1d) consistent with mTOR induction of IRE1α-XBP1 function in liver, other organs and cell types22, 23, 24. Thus, IRE1α-XBP1 induction in NK cells is at least partially driven by both STAT4 and mTOR signaling pathways.
The UPR also activates transcription factor Chop (encoded by Ddit3), a canonical ER stress marker that mediates ER stress responses. Notably, in contrast to MCMV activation of IRE1α/XBP1 in NK cells, MCMV infection repressed Ddit3 expression at day 1.5 PI compared to naïve NK cells (Fig. 1b, Supplementary Fig. 1a) as did stimulation with IL-12 and IL-18 in vitro (Fig. 1d, Supplementary Fig. 1b). In contrast, NK cells treated with the pharmacologic ER stress inducer tunicamycin increased both Ddit3 and IRE1α-XBP1 activation. (Supplementary Fig. 2a). Hence, viral infection-driven NK cell activation selectively induces a limited or “non-canonical” UPR restricted to the IRE1α-XBP1 branch.
Intrinsic requirement for IRE1α in NK cell antiviral immunity
To determine whether IRE1α-XBP1 activation in NK cells contributes to host protection against lethal viral infection, we generated mice with specific IRE1α ablation in NK cells (denoted as IRE1NK, and IRE1α-deficient NK cells henceforth denoted as IRE1NK cells, Supplementary Fig. 2a–f). MCMV-infected IRE1NK were more susceptible to MCMV infection, with significantly increased viral titers and somewhat reduced overall survival (Fig. 2a,b) than littermate control (IRE1f/f) mice (Supplementary Fig. 2a–d,f). These data demonstrate that IRE1α is required for NK cell-mediated antiviral immunity.
Intrinsic requirement for IRE1α-XBP1 in NK cell expansion
Adaptive immune features of NK cells can contribute towards host protection against viral infection25. Ly49H-expressing NK cells recognize MCMV infected cells and undergo a proliferative burst to enlarge the overall pool of effector cells, followed by a contraction of activated NK cells to form a long-lived population of memory NK cells3. To identify whether IRE1α promotes antiviral immunity of NK cells by regulating their adaptive traits, we co-transferred equal numbers of Ly49H+ IRE1NK and CD45.1 congenic wild-type (WT) NK cells into Ly49H-deficient hosts and evaluated the ability of the transferred Ly49H+ cells to expand following MCMV infection (Fig. 2c and Supplementary Fig. 3a). Whereas the transferred WT NK cells expanded robustly by day 7 PI, their IRE1NK counterparts were dramatically reduced (Fig. 2d,e). Moreover, the relative ratio of WT to IRE1NK cells remained elevated throughout the time course, ruling out a delayed expansion of the IRE1NK population (Fig. 2e).
Reduction of IRE1NK cells in peripheral blood was not due to selective aberrant trafficking, as similar deficiencies were found in spleen, liver, lung, BM and lymph nodes (Fig. 2f). A similar expansion defect was observed in NK cells lacking IRE1α’s downstream substrate XBP1 (mice termed XBP1NK, Fig. 2g, h and Supplementary Fig. 2b–d). Transcriptomics revealed no change in expression of well-defined IRE1-dependent decay of mRNA (‘RIDD’) target genes (Supplementary Fig. 4), indicating that IRE1α functions primarily via controlling its major substrate XBP1 rather than by RIDD. In contrast to the expansion defect, NK cell activation, cytokine production and cytotoxicity on a per cell basis was unaffected by deletion of IRE1α (Supplementary Fig. 3b, c). Levels of Akt-mTOR signaling were unchanged in IRE1NK cells consistent with the placement of IRE1α-XBP1 downstream of mTOR (Supplementary Fig. 3d). We conclude that there is an essential cell-intrinsic function for IRE1α-XBP1 to promote clonal expansion of NK cells during viral infection.
Control of NK cell proliferation but not survival by IRE1α-XBP1
To determine whether IRE1α-XBP1 controls proliferation or survival/apoptosis in activated NK cells, we co-transferred Ly49H+ IRE1NK and CD45.1 congenic WT NK cells labeled with a cell proliferation tracing dye into MCMV-infected Ly49H-deficient recipients and confirmed that IRE1α deficiency markedly impaired MCMV-driven proliferation in vivo (Fig. 3a–c). Diminished proliferative capacity of IRE1NK Ly49H+ cells was corroborated by their reduced ability to incorporate EdU (5-ethynyl-2’-deoxyuridine) (Fig. 3d). IRE1NK cells did not exhibit enhanced apoptosis, as determined by Annexin V or activated caspases staining (Fig. 3e, f). Hence IRE1α is required for the proliferation but not survival of activated NK cells during viral infection.
Because IRE1α activation is induced to a similar extent in both antigen-specific Ly49H+ and bystander Ly49H− 26 NK cells during MCMV infection (Fig. 1c), it was possible that IRE1α could regulate cytokine-driven NK cell proliferation independently of Ly49H-m157 engagement. Indeed, even adoptively transferred Ly49H− NK IRE1NK cells showed impaired proliferation (Fig. 3c), albeit to a lesser extent than MCMV infected Ly49H+ NK cells.
IRE1α-XBP1 drives NK cell homeostatic proliferation
Robust NK cell expansion can also be triggered by lymphopenia (homeostatic proliferation)27. Co-transfer of equal numbers of IRE1NK and CD45.1 congenic WT splenic NK cells into lymphocyte-deficient (recipient Rag2−/− Il2rg−/− mice) hosts (Fig. 4a) revealed that IRE1α deficiency leads to ~2.5 fold lower homeostatically-driven NK cells (Fig. 4b) due to decreased proliferation (Fig. 4c). Thus, although IRE1α-XBP1 may be dispensable for NK cell development and basal homeostasis at steady state (Supplementary Fig. 2b–f), IRE1NK cells exhibit dramatically compromised homeostatic proliferation during lymphopenia.
IL-2 and IL-15, cytokines of the common gamma chain family, expand28, 29 and induce XBP1 splicing in NK cells ex vivo from mouse spleen, BM and human PBMCs (Fig. 1e, f). A 1:1 co-culture of IRE1NK and CD45.1 congenic WT NK cells with IL-2 and IL-15 revealed that IRE1NK cells were continually outcompeted by WT cells (Fig. 4d). Interestingly, IRE1NK cells did not ‘blast’ (a common characteristic of proliferating cells) as much as their WT counterparts (Fig. 4e). and proliferated less well than IRE1f/f littermate NK cells cultured with IL-2 and IL-15 (Fig. 4f, g) although they expressed normal levels of these cytokine receptors (Supplementary Fig. 2f), ruling out hyposensitivity to cytokine signaling as the reason for their reduced proliferation. Additionally, pharmaceutical blockade of IRE1α activity using 4μ8C30 small molecule inhibitor of the IRE1α RNase domain – led to significant reduction in proliferation of primary human NK cells cultured in IL-2 and IL-15 (Fig. 4h, Supplementary Fig. 4b). Interestingly, 4μ8C primarily hindered the proliferation of CD56bright NK cells (Fig. 4i), a subset of human NK cells with higher proliferative capacity compared to CD56dim counterparts29 and consistent with a more robust induction of XBP1s protein in CD56bright NK cell subsets (Fig. 4j). Thus, IRE1α-XBP1’s function in growth and proliferation is conserved between mouse and human NK cells..
IRE1α-XBP1 promotes oxidative phosphorylation in NK cells
To identify key cellular processes and molecular mediators downstream of IRE1α-XBP1 in proliferating NK cells, we generated mixed IRE1NK (CD45.2): WT (CD45.1) BM chimera mice31 (Supplementary Fig. 2e) and performed whole genome transcriptome analysis on IRE1NK and WT Ly49H+ NK cells harvested at day 0, 1.5, and 7 PI. IRE1NK cells showed an overall transcriptome profile similar to their WT counterparts prior to infection, with 76 genes differentially expressed at day 0 (Fig. 5a). Importantly, the gene expression profile of IRE1NK cells at day 1.5 PI was markedly distinct from WT NK cells, with 897 genes differentially expressed (Fig. 5a).
Activated immune cells undergo robust metabolic responses, key for regulating immune cell function and long-term fitness32. Activation of NK cell oxidative phosphorylation (OXPHOS) and glycolysis is essential for robust NK cell responses33, 34. Interestingly, OXPHOS was highlighted as the top downregulated cellular process in IRE1NK cells by GO analysis (Fig. 5b, c). Metabolic flux analysis on IL-12 and IL-18 activated human primary NK cells showed significantly decreased mRNA levels of OXPHOS signature and reduced maximal respiration rate upon blockade of IRE1α activity with 4μ8C (Fig. 5d). Consistent with this, electron microscopy revealed disrupted mitochondrial morphology on sorting-purified IRE1NK cells during MCMV infection (Supplementary Fig. 5). Collectively, the molecular, cellular and metabolic evidence suggests a critical role of IRE1α-XBP1 in supporting OXPHOS in NK cells.
XBP1 promotes NK cell proliferation in part by direct transactivation of c-Myc expression
We used the QIAGEN Ingenuity Pathway Analysis platform to identify key molecular mediators of proliferation in IRE1NK cells. Both XBP1 (top 1, p = 2.66−38) and IRE1α (gene name Ern1, top6, p = 6.23−8) were among the top upstream regulators in IRE1NK cells providing validation (Supplementary Fig. 6a). Surprisingly, reduction of transcription factor Myc (N-Myc, top2, p = 3.93−28; c-Myc, top 5, p = 4.75−8) in IRE1NK vs WT NK cells was predicted and the Myc signaling pathway was also identified by GO analysis (Fig. 6a) Further, we observed significantly reduced expression of Myc targets that control cell proliferation or processes highly related to cell growth in IRE1NK cells (Fig. 6b). Although Myc controls a range of biological processes including cell growth, protein synthesis and cell metabolism35, 36, its importance in NK cell responses is less well studied33. Interestingly, although there is significant overlap between IRE1α-regulated and Myc-regulated genes identified in MCMV-primed NK cells (Supplementary Fig. 6b), Myc has not previously been described as a target of IRE1α-XBP1.
We investigated the kinetics of Myc expression in differentiating NK cells, and its potential correlation with IRE1α-XBP1 activation. N-Myc was undetectable in NK cells at all time points following MCMV infection, whereas basal levels of c-Myc mRNA were observed in naïve NK cells (Supplemental Fig. 6c). No appreciable expression of c-Myc protein was evident at baseline (Supplemental Fig. 6c), attributed perhaps to c-Myc protein instability secondary to multiple degradation mechanisms37. Following MCMV infection, NK cells robustly upregulated c-Myc mRNA, and protein levels were now easily detectable (Supplemental Fig. 6c). Furthermore, in MCMV-infected ERAI mice, levels of c-Myc protein correlated with IRE1α activity (Supplemental Fig. 6c). These data suggest that engagement of both the IRE1α-XBP1 UPR and c-Myc are important during NK cell priming.
To test whether IRE1α-XBP1 is required for c-Myc induction, we measured c-Myc expression in IRE1NK cells. The basal levels of c-Myc mRNA and protein were unaffected by the absence of IRE1α in naïve NK cells (Supplemental Fig. 6d). During MCMV infection, c-Myc transcript and protein levels were significantly diminished in IRE1NK cells compared to WT counterparts (Fig. 6c–e). Furthermore, suboptimal upregulation of canonical c-Myc targeting genes was observed in IRE1NK cells (Supplementary Fig. 6e). Consistent with the notion that c-Myc acts as a transcriptional repressor of the cell cycle checkpoint protein p2138, mRNA levels of Cdkn2a were significantly upregulated by depletion of IRE1α (Supplemental Fig. 6e). Because c-Myc expression was not completely eradicated in the absence of IRE1α, complementary upstream mechanisms likely exist (i.e. mTOR, Supplementary Fig. 6h). nduction of the c-Myc protein during homeostatic proliferation was also impaired in IRE1NK cells (Supplementary Fig. 6i). Together, these results indicate that IRE1α-XBP1 activation regulates c-Myc expression in NK cells.
XBP1s acts as a versatile transcription factor that facilitates adaptation to ER stress by regulating distinct sets of target genes involved in protein folding and quality control9. Putative XBP1-binding sites39 were found in both human and mouse c-Myc promoter regions (Supplementary Fig. 6f). Chromatin immunoprecipitation assays performed on a mouse NK/ILC cell line (MNK-1) phenotypically and functionally similar to primary mouse NK cells40, and on two human NK cell lines derived from patients with aggressive NK leukemia/lymphoma (NKL41 and KHYG-142) revealed a robust enrichment of XBP1 binding to the c-Myc promoter region (Fig. 6f),. We also utilized a c-Myc reporter mouse strain (referred to as ‘MycGFP) to assess the extent to which c-Myc promoter activation is dependent on IRE1α. MycGFP NK cells were stimulated in vitro in the presence or absence of 4μ8C. Whereas c-Myc reporter GFP was induced upon treatment with either IL-15, or IL-12 and IL-18 in DMSO-treated control groups, GFP induction was largely impeded by IRE1α blockade (Fig. 6g). Our data demonstratet that IRE1α positively regulates c-Myc transcription in activated NK cells by direct binding of XBP1 to the c-Myc promoter.
To evaluate c-Myc function in the expansion of activated NK cells, we generated mice that harbor NK cells with haploinsufficiency in c-Myc (referred to as MycNK, Supplemental Fig. 7). If IRE1α-XBP1 regulates NK cell expansion primarily via its impact on c-Myc, we would predict that MycNK cells would show a similar defect to IRE1NK cells. Indeed, when equal numbers of MycNK and CD45.1 congenic WT Ly49H+ NK cells were co-transferred into Ly49H-deficient recipients infected with MCMV, WT NK cells outcompeted MycNK cells during the entire course of the response (Fig. 5h). Thus, c-Myc deficiency impairs clonal expansion of NK cells in a manner similar to NK cells lacking IRE1α or XBP1.
To formally test the hypothesis that IRE1α-XBP1 functions at least in part by regulating c-Myc in activated NK cells to facilitate their proliferation, we generated mice that harbor NK cells with overexpression of c-Myc (Mycfsf/+ Ncr1Cre+, referred to as MycOE). Levels of c-Myc were moderately elevated by 2-fold in MycOE NK cells compared to littermate controls (Fig. 7a), although this conferred no appreciable advantage in proliferation during in vitro expansion (Fig. 7b–e). We posited that if c-Myc is a primary mediator of IRE1α-XBP1 activity, then overexpression of c-Myc should restore NK cell proliferation when IRE1α is pharmaceutically blocked or genetically depleted. Indeed, we found that during in vitro expansion, NK cells from WT littermates (Mycfsf/+ Ncr1Cre-) had impaired proliferation after treatment with 4μ8C; however, NK cells from MycOE mice were largely rescued from this defect and proliferated to a comparable extent as in the DMSO-treated control group (Fig. 7b). Thus, the requirement of IRE1α-XBP1 for NK cell proliferation can be partially bypassed by moderate overexpression of c-Myc. We further crossed MycOE with IRE1NK mice to overexpress c-Myc in NK cells with genetic deletion of IRE1α. The delayed proliferation observed in IRE1NK cells was significantly alleviated by restoration of c-Myc via genetic overexpression (Fig. 7c–e). Collectively, these findings reveal a novel XBP1-Myc axis that regulates NK cell expansion during infection and homeostatic proliferation.
Requirement for IRE1α/XBP1 in NK cell-mediated antitumor immunity
In addition to antiviral responses, NK cells play an essential role in the host antitumor response43, 44, 45, 46. The experimental intravenous B16 melanoma model provides a potent functional measurement of NK cell-mediated antitumor protection. Thus, we introduced B16F10 tumor cells intravenously to IRE1NK and IRE1f/f littermates. While the majority of IRE1f/f mice were protected from lung tumor colonization at day 10 after inoculation, IRE1NK mice exhibited significantly more melanoma nodules (Fig. 8a) and micro-metastatic lesions (Fig. 8b). Survival of melanoma-bearing IRE1NK mice was significantly decreased and inversely correlated with the increased tumor burden (Fig. 8c). Similar to IRE1NK mice, XBP1NK mice also failed to control melanoma growth (Fig. 8d), consistent with a role for XBP1 downstream of IRE1α in NK cells. Similar to the defect in expansion in IRE1NK cells during viral infection, the lungs from tumor-inoculated IRE1NK mice displayed a 50–70% reduction in percentages and absolute numbers of infiltrating NK cells compared to IRE1f/f littermates lungs (Fig. 8e), and significantly decreased levels of proliferation marker Ki-67 (Fig. 8f) and transcription factor c-Myc (Fig. 8g). Of note, the decreased number of NK cells and decreased levels of Ki-67 and c-Myc (Fig. 8e–g) were only apparent following tumor injection, as no changes were detected in tumor-free animals (Supplementary Figs. 8a, and 2b). These findings demonstrate that the IRE1α-XBP1 pathway is required for optimal NK cell expansion that facilitate host protection against NK cell-sensitive cancer.
Discussion
We demonstrate that the IRE1α-XBP1 branch of the ER Stress Response is a previously unrecognized key signaling pathway in NK cells essential for their function in viral infection, homeostatic proliferation and cancer. NK cells lacking either the upstream sensor IRE1α or its downstream substrate XBP1 fail to expand in response to MCMV infection, lymphopenia or melanoma. Mechanistically, IRE1α-XBP1 directly regulate c-Myc signaling and mitochondrial respiration to control NK cell proliferation. Hence the mammalian immune system has evolved a mechanism to incorporate a common cellular stress process to enhance antiviral immunity. The fact that this conserved pathway is co-opted by pathogens to ensure their survival in host cells47 provides intriguing evidence for host-virus coevolution.
The host immune system controls viral infection primarily through early activation of innate NK cells followed by adaptive responses by CD8+ cytotoxic T cells. NK cells recognize infected cells without prior sensitization to counteract infection by initiating effector functions including cytotoxicity, cytokine production, and proliferation7. However, less is known about the intrinsic regulators that control their function or expansion5, 6. Here we identify the ER Stress Response as an essential driver of NK cell expansion in vivo during MCMV infection. CMV is a leading cause of birth defects in newborns and a dangerous pathogen in immunocompromised individuals (including cancer and transplant patients). No effective vaccine against CMV is available48. We demonstrate that human primary NK cells activate IRE1α-XBP1 in response to pro-inflammatory cytokines present during CMV infection and rely on this pathway for optimal ex vivo expansion. IRE1α activation may help develop new vaccine strategies for host control of pathogens by strengthening NK cell responses.
In patients and animal models, impaired NK cell function is associated not only with recurring viral infection, but also with an increased incidence of various malignancies7, 49, 50. NK cells whose “primed” cytotoxicity is elicited by activating receptor signaling51, 52, and their ability to recruit other immune cells into the tumor microenvironment46 play a pivotal role in cancer immunosurveillance. The failure to control melanoma lung metastasis is associated with impaired expansion of the NK cell compartment, providing the first in vivo indication that IRE1α-driven NK cell expansion is critical for tumor immunosurveillance. The deficiency in NK cell expansion correlates with reduced recruitment of type 1 conventional dendritic cells and CD8+ T cells consistent with the notion that NK cells not only function as direct killers but also potentiate an antitumor microenvironment46. Future studies should investigate whether IRE1α-XBP1 represents the major signaling pathway that drives NK cell expansion in additional malignancies and to assess whether and to what extent other known regulators of NK cell expansion such as Zbtb325, Runx1/36 and IRF853 also contribute to halting tumor growth.
NK cells have recently attracted attention for their potential use as immune-based therapies51, 52, 54, 55, 56, best documented in the setting of hematopoietic cell transplantation (HCT) in patients with leukemia43. Alloantigen-specific NK cells can benefit such patients by providing graft-versus-tumor function while preventing T cell-mediated graft-versus-host disease44. Leukemia patients undergoing immunoablative treatments before HCT often become susceptible to CMV48, highlighting the key role of NK cells in eliminating both tumor and virus. In addition, NK cells are the first lymphocytes to reconstitute the BM of patients following HCT43. However, the magnitude of and mechanism underlying NK cell homeostatic proliferation in the setting of lymphopenia are poorly understood3, 27, 57. We demonstrate that -although unnecessary for normal NK cell expansion during development - IRE1α-XBP1 is indispensable for the homeostatic proliferation of mature NK cells in vivo in response to lymphopenia-induced stress, and for IL-15-driven homeostatic-like proliferation of human primary NK cells largely provided by the CD56bright highly proliferative subset. Our findings provide justification for incorporating NK cells into adoptive cellular immunotherapies to target cancer in patients following immunoablative treatment7, 50, and suggest that pre-treatment of NK cells with a combination of cytokines (for instance, IL-15, IL-12 and IL-18 that are now being used in the clinic) might further increase the therapeutic efficiency of adoptive NK cell transfer. As this paper was being revised after peer review, it was reported that XBP1 is important in vitro for IL-15-driven human NK cell function58. By using newly-engineered genetic mouse models, our study demenstrates an essential in vivo requirement of the IRE1α-XBP1 pathway in NK cell-mediated antitumor and antiviral protection, and identifies novel cellular and molecular mechanisms of action not previously reported. Future studies should test the feasibility and efficacy of utilizing genetically manipulated IRE1α-XBP1-overexpressing “adaptive” NK cells to treat a variety of liquid or solid malignancies.
In sharp contrast to its protective role in activating NK cell immunity against CMV and melanoma described herein, overactive IRE1α-XBP1 in dendritic cells17 and CD4+ T cells16 can cripple antitumor immune responses against ovarian cancer. The highly pleiotrophic functions of this pathway and the unique molecular mechanisms that govern those functions in different organs and cell types may explain the disparate effects of overactivation or silencing of this UPR branch. For example, the upstream pathways of IRE1-XBP1 vary among cell types. In macrophages, activation is induced through Toll-like receptor signaling while in NK cells, as shown here, both IL-12-STAT4 and mTOR act as critical upstream regulators of IRE1α-XBP1 under infectious/inflammatory conditions. Indeed mTOR is also situated downstream of IL-1528, 59 and IL-1233 and is essential for NK cell development and antitumor responses60. A recent publication demonstrated that mTOR drives IRE1α-XBP1 activation in liver in response to food perception22, as well as in lung TH2 cells and fibroblasts in fibrotic disease23, 24.
We also report the unexpected finding that XBP1 directly controls the transcription of c-Myc and its downstream target genes in NK cells. This may appear paradoxical in light of the recent finding that the oncogene c-Myc acts upstream of the IRE1α-XBP1 pathway to drive tumorigenesis61. However, this discrepancy may be explained by the differing functions and downstream consequences of IRE1α signaling in normal versus transformed cells. One clear difference between transformed versus normal cells is that the former are addicted to constitutively activated c-Myc while the latter display negligible levels of c-Myc at baseline, requiring external stimuli to be induced. Although c-Myc is among the best-studied oncogenes and tumor-intrinsic c-Myc activity can program an immune suppressive microenvironment obligatory for tumor progression62, its function in immune cells is less understood63. c-Myc is essential for metabolic responses in IL-2 and IL-12-activated mouse NK cells ex vivo33. In human NK cells ex vivo, c-MYC functions downstream of IL-15 to enhance effector function64. Here, we provide the first in vivo evidence of an essential role for c-Myc acting downstream of IRE1α-XBP1 to drive NK cell expansion during MCMV infection, lymphopenia and antitumor immunity. Indeed c-MYC levels were noted to be significantly decreased in NK cells from cancer patients compared to healthy donors65, suggesting a protective role of c-MYC in NK cell-mediated antitumor immunity in patients.
Finally, our finding that IRE1α supports mitochondrial function and morphology in NK cells was unexpected given that we have recently described that depletion of IRE1α-XBP1 helps to restore the metabolic fitness of T cells in ovarian cancer16. Intriguingly, 25 of the 46 differentially expressed OXPHOS genes in IRE1α-deficient NK cells are known to be regulated by c-Myc35. Thus, the dysregulated metabolism observed here may be in part a consequence of impaired Myc signaling33 which is in turn at least partly responsible for the failure of IRE1α-deficient NK cells to expand appropriately in vivo in response to viral infection, lymphopenia and malignancy.
In conclusion, our study reveals a novel and essential function for the ER stress sensor IRE1α and its downstream substrate XBP1 to drive NK cell expansion that results in effective responses against viral infection and tumors. IRE1α activation is situated downstream of both the Stat4 and mTOR signaling pathways and upstream of c-Myc. Like c-Myc, IRE1α-XBP1 also promotes OXPHOS in NK cells. We have identified a novel IRE1α-XBP1-cMyc axis in NK cell immunity, required for host protection against infection and cancer.
Methods
Mice
Mice were bred at Dana-Farber Cancer Institute in accordance with the guidelines of the Institutional Animal Care and Use Committee (IACUC). The following strains were used in this study, all on the C57BL/6 genetic background: C57BL6/J (CD45.2+; The Jackson Laboratory), B6.SJL (CD45.1+; The Jackson Laboratory), Rag2−/−Il2rg−/−(Taconic), Klra8−/− (Ly49H-deficient)66, IRE1f/f67, XBP1f/f11, Mycf/f (a generous gift from Dr. Ruoning Wang), Ncr1iCre (a generous gift from Dr. Eric Vivier), ERAI (ER stress-activated indicator)19, and GFP-c-Myc KI (here called MycGFP, the Jackson Laboratory). Mycfsf/fsf on mixed B6.126 background was a generous gift from Dr. Rosaline Sears. IRE1f/f Ncr1iCre/WT (here called IRE1NK), XBP1f/f Ncr1iCre/WT (XBP1NK), Mycf/+ Ncr1iCre/WT (MycNK), Mycfsf/+ Ncr1iCre/WT (MycOE) and Mycfsf/+ IRE1f/f Ncr1iCre/WT (MycOE IRE1NK) animals and littermate controls were generated by breeding at Dana-Farber Cancer Institute. Recipient Klra8−/− (CD45.1+CD45.2+) mice were generated by crossing Klra8−/− (CD45.2+) to B6.SJL (CD45.1+). Experiments were conducted using age- and gender-matched mice in accordance with approved institutional protocols.
Virus infection
MCMV (Smith strain) was passaged serially through BALB/c hosts two times, then viral stocks were prepared by using a dounce homogenizer to dissociate the salivary glands of infected mice 3 weeks after infection. For direct infection of C57BL/6 or ERAI mice, 7.5 × 103 plaque-forming units (PFU) of MCMV was injected by i.p.. For the adoptive transfer studies, Ly49-deficient mice were infected by i.p. injection of 7.5 × 102 PFU of MCMV one day after receiving approximately 2 × 105 Ly49H+ NK cells by intravenous (i.v.) injection.
NK cell enrichment and adoptive transfer
Enrichment of NK cells was performed using the magnetic bead-based negative selection kit (Miltenyi Biotec) per the manufacturer’s protocol (typically 50–70% purify) before the flow cytometric analysis of the transferred NK cells in the spleen of recipients at early time points after infection of MCMV in some experiments (Fig. 3c–f, Fig. 6e). As previously described31, for experiments involving co-transfer of CD45.1+ and CD45.2+ populations, equal numbers of Ly49H+ NK cells (normalized on KLRG1Lo population) from each population were co-transferred together by intravenous injection into the tail vein of adult Ly49H-deficient recipients 1 d before infection with MCMV, or equal numbers of NK cells from each population were co-transferred together into adult Rag2−/−xIl2g−/− recipients. WT and KO NK cells are transferred together into the same mouse and the numbers of cells are the same. Thus, the percentages shown are directly correlated with the absolute number of cells. In some experiments, cells were labeled before transfer with CellTrace Violet (CTV; Invitrogen) to trace cell proliferation.
Flow cytometry and cell sorting
Cell surface staining of single-cell suspensions was performed using the following fluorophore-conjugated antibodies (purchased from BD Biosciences, eBioscience, BioLegend, Tonbo Biosciences, Cell Signaling Technology, and R&D Systems): NK1.1 (PK136), KLRG1 (2F1), Ly49H (3D10), CD45.1 (A20), CD45.2 (104), CD8α (53–6.7), TCRβ (H57–597), CD3ε (145–2C11), F4/80 (BM8), CD19 (eBio1D3), Ki-67 (SolA15), c-Myc (D84C12), and XBP1s (Q3–695). Unless otherwise indicated, NK cells were defined as CD19−CD3− TCRβ− F4/80− NK1.1+ cells. All analysis was performed on viable populations by exclusion of the dead cells using a fixable viability dye (Thermo Fisher). For the evaluation of NK cell apoptosis, Pan-caspase (FLICA) staining was carried out using the FAM FLICA in vitro Poly Caspase Kit (Immunochemistry Technologies) per the manufacturer’s protocol; Annexin V staining was carried out using the Invitrogen eBioscience Annexin V Apoptosis Detection Kit per the manufacturer’s instructions. For the evaluation of NK cell proliferation, cells were labeled with 5 uM CTV (Invitrogen) according to the manufacturer’s protocol before transfer into recipient mice; intracellular staining of Ki-67, c-Myc and XBP1s was performed by fixing and permeabilizing with the Foxp3/Transcription Factor Staining Kit (eBioscience); for the EdU labeling experiments, mice were subjected to i.p. injection of 1.5 mg of EdU in PBS on days 3.5 after infection with MCMV, and EdU labeling was determined ~16 h later using the Click-iT EdU Assay Kit (Thermo Fisher Scientific). Flow cytometry was performed on the LSR II (BD Biosciences), LSR Fortessa X-20 and CytoFLEX LX (Beckman Coulter). Cell sorting was performed on Aria II cytometers (BD Biosciences). For experiments involving qRT-PCR, cell populations were sorted to >95% purity. Data were analyzed with FlowJo software (Tree Star).
Ex vivo stimulation of primary mouse and human NK cells
For Fig. 1e, ~2 × 105 NK cells were stimulated for 16 hr in complete RPMI containing 10% FBS with recombinant mouse IL-2 (10 μg/ml; BioLegend), either in the presence or absence of IL-12 (20 ng/ml; Peprotech) and IL-18 (10 ng/ml; R&D Systems), or IL-15/15R (100 ng/ml, eBioscience. For ex vivo NK cell expansion experiments, 1.5 × 105 to 2.5 × 105 purified NK cells were stimulated in RPMI supplemented with 10% FBS, penicillin-streptomycin, L-glutamine and β-mercaptoethanol, and with recombinant mouse IL-2 (10 ng/ml; R&D Systems) and IL-15/15R (40 ng/ml; eBioscience) re-applied every other day. In some assays indicated, 5 uM of 4μ8C (IRE1α inhibitor; Sigma Aldrich) was added to the culture and DMSO was used as the control treatment.
Peripheral blood mononuclear cells (PBMCs) from healthy donors (New York Blood Center) were isolated by density gradient centrifugation with Ficoll (GE Healthcare). NK cells were enriched by negative selection using the human NK Cell Isolation Kit (Miltenyi Biotec; >90% purity). For ex vivo stimulation in Fig. 1f, ~106 sorted primary human NK cells were stimulated for 16 hr in complete RPMI containing 10% FBS with recombinant human IL-2 (10 μg/ml; Sigma Aldrich), in combination with either IL-12 (20 ng/ml; StemCell) and IL-18 (10 ng/ml; Thermo Figher Scientific), or IL-15 (40 ng/ml, StemCell). For human NK ex vivo expansion in Fig. 4h, i, recombinant human IL-2 (100U/ml, Roche) and IL-15 (10 ng/ml, StemCell) was applied. Fresh medium and cytokines were added on alternating days of culture. In some assays indicated, 4μ8C (IRE1α inhibitor, 5 μM or lower) was added to the culture and DMSO was used as the control treatment. For flow cytometric analysis or cell sorting, human NK cells were defined as CD3− CD56+.
NK cell lines and culture conditions
MNK-1 cells40 were cultured in DMEM supplemented with 10% FBS, 2 mM GlutaMAX, 1 mM sodium pyruvate, 55 μM 2-mercaptoethanol, 10 mM HEPES, 50 μg/ml gentamicin, 100 U/ml penicillin, and 100 μg/ml streptomycin (all from Corning or Gibco Life Technologies, except FBS from Atlanta Biologicals). To facilitate robust survival, 10 ng/ml recombinant mouse IL-2 (BioLegend) were re-applied daily. NKL cells41 and KHYG-1 cells42 were cultured in RPMI 1640 medium supplemented with 2 mM L-glutamine, 1 mM sodium pyruvate, 100 U/ml penicillin, 100 μg/ml streptomycin and 7.5% FCS plus 7.5% new born calf serum (all from Corning or Gibco Life Technologies, except FBS from Atlanta Biologicals). The cell concentration was maintained at ~5 × 105 /ml in the presence of 50 U/ml of recombinant human IL-2 (Amgen). All cell lines routinely tested negative for mycoplasma contamination using the Mycoplasma Detection Kit (Lonza) and maintained in medium containing plasmocin (Invivogen) for prophylactic purpose.
Quantitative real time PCR
Cells were pre-enriched (Miltenyi Biotec) and then sorted to >95% purity and RNA was isolated using the RNeasy RNA purification mini (for 5 × 105 cells or beyond) or micro (for <5 × 105 cells) kit (Qiagen) according to the manufacturer’s protocol. From purified RNA, cDNA was synthesized using the reverse-transcriptase kit qScript cDNA synthesis kit (Quanta Biosciences). Quantitative real time PCR (qRT-PCR) was performed in triplicates in a 384-well plate using Taqman Probes (Themo Fisher) or SYBR Green (Quanta Biosciences) -based detection on an ABI QuantStudio 6 Flex qRT-PCR machine. For experiments in Fig. 6c, CellDirect One-Step qRT-PCR Kit with Rox (Life Technologies) was used according to the manufacturer’s instruction. For the analysis of mRNA abundance, the derived values were normalized to those of β-Actin (β-ACTIN for human materials). Primer sequences are in Supplementary Table 1 and Taqman probe information is in Supplementary Table 2.
RNA-seq analysis
Mixed IRE1NK (CD45.2): WT (CD45.1) BM chimeric mice were generated as previously described31. Briefly, host C57BL/6 × B6.SJL animals (CD45.1+CD45.2+) were lethally irradiated with 900 grays of radiation and reconstituted with a 1:1 mixture of BM cells from B6.SJL WT (CD45.1+) and knockout or transgenic donor (CD45.2+) mice, co-injected with anti-NK1.1 (clone PK136) to deplete any residual donor or host mature NK cells. CD45.1+CD45.2+ host NK cells were excluded from all analyses.
Mice were infected with 7.5 × 103 PFU of MCMV by i.p. injection and spleens were harvested at day 0, 1.5 and 7 after infection. Total RNA from sorting-purified Ly49H+ WT or IRE1NK NK cells was extracted with the RNeasy RNA purification micro kit (Qiagen) according to the manufacturer’s instructions. Poly(A)+ RNA-seq libraries (pair-end 100) were prepared with an Illumina TruSeq RNA Sample Preparation Kit V2 according to the manufacturer’s instructions (Illumina). Sequence data from Illumina’s HiSeq4000 sequencer were ‘de-multiplexed’ to generate FASTQ files for each sample with Illumina’s CASAVA pipeline (version 1.8.2).
The reads that passed Illumina’s quality and purity filter (> 90%) were aligned to the mouse genome (Illumina iGenomes mm9 build) with tophat268 with default parameters. The resulting SAM alignment files were then converted to the BAM file format, then were sorted and indexed with SAMtools (version 0.1.14). The HTSeq69 was used for summarization of the aligned sequence reads into counts. Differential gene expression was analyzed with the DESeq270. Transcripts with adjusted P value < 0.01 were considered differentially expressed between genotypes (wild-type and IRE1NK) and infection times (day 0, 1.5, 7). Gene ontology analysis of differentially expressed genes was performed in GSEA (gene sets enrichment analysis)71. Pathway analysis of predicted upstream regulators was performed with Ingenuity Pathway Analysis software (Qiagen). For heat maps, normalized data from DESeq RNA-seq analyses were exported, manually curated for specific genes in each category, and the gene-expression z-score was visualized with the Heatmap.2 function within the gplots R library.
Chromatin immunoprecipitation (CHIP)
MNK-1, NKL and KHYG-1 cells were crosslinked with 1% formaldehyde for 10 min at room temperature. Reaction was quenched with 125mM glycine. CHIP was performed as previously described72 with XBP1 antibody (Cat. 619502, Biolegend), or normal rabbit IgG control (Cat. #2729, CST). The sequences of all primers are listed in Supplementary Table 3.
Immunoblots
Sorted purified NK cells (purity>97%) from WT: IRE1NK mixed BM chimeras day 1.5 PI were collected directly into 20% TCA (trichloroacetic acid) buffer. After sorting, the concentration of TCA was adjusted to 10% before incubation on ice for 30 min to precipitate proteins. The homogenates were centrifuged at 13, 000 r.p.m. for 10 min at 4°C, and the supernatant was carefully removed. The pellets with concentrated proteins were washed twice with acetone, and then solubilized in solubilization buffer (containing 9M Urea, 2% Triton X-100, and 1% DTT) mixed with LDS buffer (Invitrogen). The samples were heated at 70°C for 10min and protein concentrations determined using a BCA protein assay kit (Thermo Fisher Scientific). Equivalent amounts of protein for WT and KO cells from the same BM chimeras were separated by SDS–PAGE (Bis-Tris gel, Invitrogen) and transferred onto PVDF membranes following the standard protocol. Anti-cMyc (D84C12, Cell Signaling Technology), anti-β-Actin (Cell Signaling Technology) and goat anti-rabbit secondary antibodies conjugated wiih HRP (Thermo Fisher Scientific) were used. SuperSignal West Pico and Femto chemiluminescent substrates (Thermo Fisher Scientific) were used to image blots in a ChemiDoc™ MP Imaging System instrument (BioRad). Densitometric quantification was performed using Image J software (NIH).
Metabolic flux assays
Purified human primary NK cells from healthy PBMC donors were stimulated with human recombinant IL-12 and IL-18 for 16hr in complete RPMI medium, in the presence or absence of IRE1 inhibitor 4μ8C (5 μM). After incubation, NK cells were collected, thoroughly washed and then subjected to Seahorse analyses using non-buffered XF base medium containing 25 mM glucose, 2 mM L-glutamine, and 1 mM sodium Pyruvate but lacking serum. XF96 cell-culture microplates were coated with CellTak (Corning) before analysis according to the manufacturer’s instructions and washed twice with distilled water. 6 × 105 NK cells were plated and OCR measurements were analyzed on an XFe96 Extracellular Flux Analyzer (Agilent). After Basal OCR measurements were obtained, an OCR trace was recorded in response to oligomycin (1 μM), carbonyl cyanide-p-(trifluoromethoxy) phenylhydrazone (FCCP, 1 μM), and rotenone and antimycin (0.5 μM each) following the XF Cell Mito Stress test kit (Agilent). Metabolic parameters were calculated as follows: basal OCR = last rate measurement before oligomycin injection – minimum rate measurement after rotenone and antimycin injection; maximal OCR = maximum rate measurement after FCCP injection – minimum rate measurement after rotenone and antimycin injection. Typically, 3–6 technical replicates per each sample were examined. OCR values were normalized by protein input amount accordingly.
Experimental melanoma lung metastasis
B16F10 cells73 were cultured in DMEM supplemented with 10% FBS, 100 U/ml penicillin and 100 ug/ml streptomycin. 2.0 × 105 B16F10 cells were injected intravenously through the tail vein. Mice were sacrificed and lungs were harvested on day 10–20 post inoculation. Lung tumor nodules were enumerated by gross count of visible sites of disease. Micrometastatic lesions were identified using H&E stained lung sections. All pulmonary tumor measurements were performed in a blinded manner. For the flow cytometric analysis, mice received an intravenous injection of fluorescence-conjugated anti-CD45.2 three minutes prior to euthanasia to label intravascular immune cells that would be excluded from downstream analysis. Lungs were harvested from tumor-bearing mice following euthanasia by CO2 inhalation, and single-cell suspensions were obtained using a gentleMACS dissociator (Miltenyi Biotec) per manufacturer’s instructions.
Statistics
For graphs, data are shown as mean ± s.e.m. or s. d. as indicated. and, unless otherwise indicated, statistical differences were evaluated using a two-tailed unpaired Student’s t-test, assuming equal sample variance. P < 0.05 was considered significant. Graphs were produced and statistical analyses were performed using GraphPad Prism or R. Sample size was not specifically predetermined, but the number of mice used was consistent with prior experience with similar experiments.
Reporting Summary
Further information on research design is available in the Nature Research Reporting Summary linked to this article.
Data availability
The RNA-seq data were deposited in the Gene Expression Omnibus under the accession number GSE 113214. The remaining data that support the findings of this study are available from the corresponding authors upon request. Materials will be provided with material transfer agreements in place as appropriate.
Supplementary Material
Acknowledgements
We thank E. Vivier (Aix Marseille University) for NCR1iCre mice; R. Wang (Nationwide Children’s Hospital) for Mycfl/fl mice; R. Sears (Oregon Health & Science University) for Mycfsf/fsf mice; J. Ritz (Dana-Farber Cancer Institute) for the NKL cell line; C. Hurley (Georgetown University) for the KHYG-1 cell line; D. Lyden (Weill Cornell Medicine) for the B16F10 cell line; members of the Sun lab for sharing viral reagents, cytokines and flow cytometry antibodies, and for providing experimental assistance; members of the Glimcher lab – M. Raundhal, C. Lentucci, R. Xu, S. Shrestha and A. Gonzalez for technical support; S. Adoro (Case Western Reserve University) and J. Cubillos-Ruiz for critical reading of the manuscript and helpful discussion; M. Song (Weill Cornell Medicine) and the Wucherpfennig lab (Dana-Farber Cancer Institute) for insightful comments; J. Xiang (Weill Cornell Genomics and Epigenomics Core) for RNA-seq; J. McCormick (Weill Cornell Medicine) and DFCI Flow Cytometry Core for cell sorting; Specialized Histopathology Core at Brigham & Women’s Hospital for histology; L. Cohen-Gould and J. Jimenez (Microscopy and Image Analysis Core Facility at Weill Cornell Medicine) for electron microscopy; X. Liu (Metabolism and Mitochondrial Research Core, Beth Israel Deaconess Medical Center) for metabolic flux assays; D. Neuberg (Dana-Farber Cancer Institute) for advising on statistical analysis; S. Schneider (Dana-Farber Cancer Institute) for manuscript editing. L.H.G. and H.D. were supported by Institutional Funding from The Dana-Farber Cancer Institute. H.D. was also supported by a Helen Gurley Brown Presidential Initiative Award (Dana-Farber Cancer Institute). J.C.S. was supported by the Ludwig Center for Cancer Immunotherapy, the American Cancer Society, the Burroughs Wellcome Fund, and the NIH (AI100874, AI130043, and P30CA008748). N.M.A.was supported by an MSTP Grant from the NIGMS of the NIH to the Weill Cornell/Rockefeller/Sloan Kettering Tri-Institutional MD-PhD Program (T32GM007739), and by an F30 Predoctoral Fellowship from NIAID of the NIH (F30 AI136239). X. C. was supported by a NIH 1 R37 CA228304–01 grant.
Footnotes
Competing interests
L.H.G. is a former Director of Bristol-Myers Squibb and is currently on the board of directors of and holds equity in GlaxoSmithKline Pharmaceuticals and the Waters Corporation. She chairs the scientific advisory board, is a co-founder of and holds equity in Quentis Therapeutics. She also serves on the scientific advisory boards of Repare Therapeutics, Abro and Kaleido Therapeutics.
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Associated Data
This section collects any data citations, data availability statements, or supplementary materials included in this article.
Supplementary Materials
Data Availability Statement
The RNA-seq data were deposited in the Gene Expression Omnibus under the accession number GSE 113214. The remaining data that support the findings of this study are available from the corresponding authors upon request. Materials will be provided with material transfer agreements in place as appropriate.