CD8 T lymphocytes are able to eliminate nascent tumor cells through a process referred to as immunosurveillance. However, multiple inhibitory mechanisms within the tumor microenvironment have been described that impede tumor rejection by CD8 T cells, including increased signaling by inhibitory receptors. Lysophosphatidic acid (LPA) is a bioactive lysophospholipid that has been shown repeatedly to promote diverse cellular processes benefiting tumorigenesis. Accordingly, the exaggerated expression of LPA and LPA receptors is a common feature of diverse tumor cell lineages and can result in elevated systemic LPA levels. LPA is recognized by at least six distinct G protein–coupled receptors, several of which are expressed by T cells, although the precise function of LPA signaling in CD8 T-cell activation and function has not been defined. Here, we show that LPA signaling via the LPA5 receptor expressed by CD8 T cells suppresses antigen receptor signaling, cell activation, and proliferation in vitro and in vivo. Importantly, in a mouse melanoma model tumor-specific CD8 T cells that are LPA5-deficient are able to control tumor growth significantly better than wild-type tumor-specific CD8 T cells. Together, these data suggest that the production of LPA by tumors serves not only in an autocrine manner to promote tumorigenesis, but also as a mechanism to suppress adaptive immunity and highlights a potential novel target for cancer treatment. Cancer Immunol Res; 1(4); 245–55. ©2013 AACR.
The adaptive immune system is able to detect and eliminate nascent tumors through a process referred to as immune surveillance and mediated in large part by cytotoxic CD8 T cells. However, this immune response to tumor may also contribute to tumorigenesis by providing selective pressure to which tumors adapt and eventually evade eradication, a process coined cancer immunoediting (1, 2). Tumors that evade the initial T-cell response create an immunosuppressive microenvironment from which variants arise, escape immune control, and grow without restraint. The mechanisms used by tumors to either evade the initial CD8 T-cell response or promote the tumor immunosuppressive environment are not fully defined. However, the recent success of immunotherapies that interfere with tumor-derived immune suppression (3–5) has underscored the importance of identifying the mechanisms by which tumors suppress CD8 T cells to escape immune control (6, 7).
The activation of cytotoxic CD8 T cells by either foreign or tumor antigen is initiated via signals transmitted by the T-cell antigen receptor (TCR; ref. 8). TCR signaling and the subsequent function of mature T cells can be regulated in a positive or negative manner by different surface coreceptors (9). Multiple inhibitory mechanisms within the tumor microenvironment that impede tumor rejection by tumor-infiltrating lymphocytes (TIL) have been described (6, 10), including the increased signaling by CD8 T-cell inhibitory receptors, such as the well-characterized CTLA-4 molecule (11).
Lysophospholipids comprise a small family of structurally simple lipids that include sphingosine-1-phosphate (S1P) and lysophosphatidic acid (LPA), and they induce diverse biologic and pathophysiologic effects by signaling through specific G protein–coupled receptors (GPCR; refs. 12, 13). The LPA lysophospholipid is recognized by six specific GPCR, LPA1–6 (14), and although T lymphocytes are known to express several LPA receptors (13, 15–17), the immune regulatory activities of LPA are not well understood. LPA concentration in the blood of healthy individuals has been reported to vary from high nanomolar to low micromolar levels (12, 18). In vivo, production of LPA results predominantly from the activity of autotaxin (ATX; ref. 19), an extracellular lysophospholipase D originally isolated and identified from a human melanoma as an autocrine motility factor (20). Since then LPA has been found aberrantly produced in a number of different malignant cell types (21–23), resulting in significantly increased systemic levels that can reach 60 μmol/L in malignant effusions (24–26). At these elevated levels, LPA has been shown to promote tumor progression by enhancing tumor migration, survival, metastasis, angiogenesis, and therapeutic resistance (27–31).
Previously LPA has been shown to modulate the activation of different cell types (17), and in this study, we investigated if LPA could influence CD8 T-cell activation. Here, we report that CD8 T cells express the LPA5 receptor and signaling by this GPCR inhibits CD8 T-cell receptor signaling, activation, and proliferation. Furthermore, we show that tumor-specific CD8 T cells lacking LPA5 can control the progression of established tumor progression more efficiently than the LPA5-sufficient tumor-specific CD8 T cells. Thus, our findings reveal a novel role for lysophospholipid-mediated protection of tumor from adaptive immunity.
Materials and Methods
C57BL/6 (CD45.2) and CD45.1 (B6.SJL-Ptprca Pepcb/BoyJ) were obtained from The Jackson Laboratory or bred in the Biological Resource Center (BRC) at National Jewish Health (NJH; Denver, CO). CD45.1 OT-I mice (a gift from Dr. Ross Kedl, University of Colorado, Denver, CO), Lpar2−/− mice (a gift from Dr. Jerold Chun, Scripps Research Institute, La Jolla, CA), Lpar5−/− mice, TCRα−/− mice (a gift from Dr. Philippa Marrack, NJH) were bred in the BRC at NJH. LPA5−/− mice were generated as described in the Supplementary Data. All mice used were 8 to 12 weeks of age, housed under specific pathogen-free conditions, and were maintained in accordance with the regulations of the Institutional Animal Care and Use Committee.
Erythrocyte-lysed cells isolated from spleen, pooled lymph nodes, or purified CD8+ T cells were suspended at 20 × 106 cells/mL in RPMI-1640 medium supplemented with 2.5% fatty acid–free (faf) bovine serum albumin (BSA; Calbiochem) and loaded with Indo1-AM (Molecular Probes) as described in the Supplementary Data.
CD8+ T cells were isolated from the spleens and lymph nodes of wild-type mice and LPA receptor expression measured by real-time quantitative reverse transcriptase (RT-PCR) using Platinum SYBR Green quantitative PCR (qPCR) SuperMix-UDG (Invitrogen Life Technologies). The details of qPCR analysis including specific forward and reverse LPA receptor primers are provided in the Supplementary Data.
All antibodies were purchased from eBiosciences, BD Pharmingen, Biolegend, or were produced in our laboratories. Cells were stained in 2% BSA–PBS+0.1% sodium azide with blocking Fc receptor antibody (2.4G2) on ice for 20 to 30 minutes. For viability assessment, 7-amino actinomycin D (7-AAD) was added 10 minutes before data acquisition. All flow-cytometric analysis was conducted on the LSRII flow cytometer (BD) and analyzed with FlowJo v8 (TreeStar) and GraphPad Prism software (v 5.0).
LPA (C16 LPA; Avanti Polar Lipids) was solubilized to 5 mmol/L concentration in 0.1% BSA–PBS, aliquoted, and stored at −20°C. Aliquots were diluted to 1 mmol/L in RPMI-1640 medium supplemented with 2.5% faf-BSA (Calbiochem) before use. Octodecenyl thiophosphate (OTP) was generated as previously described (32), stored as a powder, and solubilized in 95% methanol to create aliquots. Virgin glass tubes and caps were sterilized by autoclave for aliquots that were stored at −20°C. The concentration was confirmed by phosphorus assay. OTP was solubilized for experimental use by sonication with FBS- or BSA-containing culture media or vehicle (2% propanediol and 1% ethanol in PBS) for in vitro or in vivo usage, respectively. For in vitro experiments, OTP was solubilized to 50 μmol/L and passed through a 0.2-μm filter for further sterilization. For in vivo experimentation, solubilized OTP was transferred to siliconized Eppendorf tubes and animals were dosed at 5 mg/kg every 8 hours.
Generation of bone marrow–derived dendritic cells
Congenic gender-matched bone marrow–derived dendritic cells (BMDC) were generated by flushing of femur and tibia and culture at 106 cells/mL in RPMI-1640 with 20 ng/mL granulocyte macrophage colony-stimulating factor (GM-CSF), 10% FBS (Omega Scientific), penicillin–streptomycin, and GlutaMAX (Invitrogen). Media was refreshed on days 3 and 5. On day 7, BMDCs were harvested from culture and stimulated with 1 ng/mL LPS for 90 minutes and pulsed with peptide for the last hour of LPS treatment. BMDCs were washed five times to remove LPS and unbound peptide before transfer.
In vitro T-cell activation and proliferation
To determine how LPA affected antigen-specific activation of CD8 T cells, OT-I splenocytes were isolated, erythrocyte lysed, and labeled with carboxyfluorescein succinimidyl ester (CFSE; Invitrogen). For all CFSE labeling, cells were suspended at 15 × 106 cells/mL in PBS and CFSE was added to a final concentration of 2 μmol/L for 10 minutes and then washed in media. Splenocytes were pulsed with 1 μmol/L of the SIIGFEKL (G4; AnaSpec, Inc.) or SIINFEKL (a gift from Philippa Marrack) peptides for 4 hours or 90 minutes, respectively, in 5% faf-BSA RPMI, then washed. Cells were cultured in 96-well plates at 2.5 × 106 cells/mL in the presence or absence of 50 μmol/L OTP that was sterile-filtered before addition to culture. Cells were enumerated by flow cytometry, and the proportion of cells proliferated was calculated by FlowJo analysis. The mean fluorescence intensity (MFI) values of activation marker expression were normalized.
To assess in vitro cytokine production, OT-I effector T cells were generated by pulsing erythrocyte-lysed OT-I splenocytes with 1 μmol/L SIINFEKL and culture with interleukin-2 (IL-2) for 5 days. On day 5 of culture, target cells (EL4 cells) were pulsed with 1 μmol/L SIINFEKL and cultured at an effector to target ratio of 0.625:1 with OT-I effector T cells for 4 hours in the presence of Brefeldin A, in the presence or absence of sterile-filtered 50 μmol/L OTP.
In vivo T-cell transfer and antigen-specific stimulation
BMDCs were generated as described earlier. One day before BMDC transfer, CD8+ T cells were purified from OT-I spleen and lymph node cells with a CD8+ T-cell enrichment kit (Miltenyi) to a purity of 95% or more, and 106 CFSE-labeled CD8+ T cells were transferred to CD45 allotype-mismatched recipient C57BL/6 mice. SIINFEKL-BMDC (106) were suspended in PBS and transferred subcutaneously in the scruff of individual recipients. On day 3 postimmunization, animals were sacrificed and draining lymph nodes (dLN; axilary, brachial, and cervical), non-draining lymph nodes (ndLN; inguinal and mesenteric), and spleen were harvested. After erythrocyte lysis, cells were counted by Z2 Coulter Particle Count and Size Analyzer (Beckman-Coulter) and 10 × 106 cells were stained for flow cytometry. Cells were suspended in fluorescence-activated cell sorting (FACS) buffer and stained with 7-AAD for viability before analysis on the BD LSRII.
B16.cOVA tumor experiments
The OVA-transfected B16 tumor cell line (B16.cOVA) was kindly provided by Dr. Ross Kedl. Cells were maintained in RPMI-1640 (Cellgro), supplemented with 10% heat-inactivated FBS (Omega Scientific), GlutaMAX, penicillin–streptomycin, minimum essential medium (MEM) nonessential amino acids, sodium pyruvate (Invitrogen), and 0.75 mg/mL G418 sulfate selection (Cellgro).
To determine how LPA5−/− OT-I T cells responded to tumor, 105 B16.cOVA cells were transferred subcutaneously into the hind leg of recipients, and 5 days later, 0.5 × 106 CFSE-labeled OT-I T cells were transferred via the tail vein. Recipients were sacrificed 5 days later and tumor and dLN were harvested. Lymphoid organs were mashed through a cell strainer (100 μm; BD), erythrocyte lysed, and counted. Tumors were digested in 0.5 mg/mL collagenase D (Fisher Scientific) and 60 U/mL DNase (Sigma-Aldrich) for 30 minutes with perturbation every 10 minutes, and then neutralized with 5 mmol/L EDTA for 5 minutes. Tumor cells were homogenized by pipette tituration and passed through a 100 μm filter, erythrocyte lysed, and counted. Tumor diameter was measured with calipers and body weight and overall appearance were assessed every 2 days. Mice were sacrificed when tumor diameter exceeded 10 mm or at 8 days after T-cell transfer, whichever came first.
All statistical analysis was conducted with GraphPad Prism software (version 5.0) using a two-tailed unpaired Student t test.
LPA inhibits T-cell antigen receptor-mediated calcium mobilization via LPA5
Both human and murine T cells express LPA receptors (15, 16), so we initially questioned whether LPA could influence TCR signaling. Specifically, we tested whether LPA could modulate TCR-induced intracellular calcium mobilization. These results revealed that 20 μmol/L of LPA potently inhibited intracellular calcium mobilization by CD8+ T cells in response to TCR stimulation achieved by antibody-mediated cross-linking of the TCR (Supplementary Fig. S1). In contrast, LPA treatment in the absence of antigen receptor stimulation did not affect intracellular calcium levels. LPA inhibition of CD8 TCR signaling was further shown to be dose dependent with a reduction of intracellular calcium levels achieved with LPA concentrations as low as 1 μmol/L (Fig. 1A), approximating the levels reported in the blood of healthy individuals (12, 18).
There are currently six validated LPA receptors: LPA1–6 (14) and qPCR analyses revealed that purified CD8 T cells expressed LPA2, LPA5, and LPA6 (Fig. 1B), consistent with previous findings (15, 16). To determine which LPA receptor(s) was responsible for LPA inhibition, we compared calcium mobilization in TCR-stimulated CD8+ T cells from LPA2−/− and LPA5−/− mice (LPA6−/− mice were not commercially available). These results revealed that although LPA inhibition was intact in the LPA2-deficient T cells, LPA inhibition was absent in LPA5-deficient T cells (Fig. 1C). These data show that LPA5 expression is required for the inhibition of TCR-mediated calcium mobilization.
LPA signaling inhibits proliferation and activation in vitro
Increased intracellular calcium concentration is an important early consequence of antigen receptor signaling that leads to the activation of distinct transcriptional programs important for T-cell activation and function (33). Thus, we next tested whether LPA suppression of TCR-induced calcium mobilization would lead to reduced CD8 T-cell activation in vitro. Initial experiments revealed that LPA was able to inhibit TCR-mediated activation of CD8+ T cells, as determined by reduced CD62L downregulation (34), however, only when T cells were stimulated with suboptimal concentration of anti-CD3 (Supplementary Fig. S2). To investigate whether LPA could inhibit antigen-specific stimulation of CD8 T cells, we used the well-characterized OT-I TCR transgenic mouse model that features CD8 T cells that express a TCR specific for the chicken ovalbumin peptide, SIINFEKL, in the context of the MHC molecule, H-2Kb (35). This system also allowed us to modulate T-cell activation with different stimulatory peptides (36). More specifically, the OT-I TCR displays relatively high affinity (KD ∼ 5 μmol/L) for the SIINFEKL ovalbumin peptide, but it also recognizes the altered peptide ligand, SIIGFEKL (G4), with relatively weaker affinity (KD ∼ 10 μmol/L; ref. 37). Thus with these peptides, we could compare the effects of LPA signaling after relatively high or low affinity TCR stimulation. Because LPA is degraded both in vitro and in vivo (38), we also used a metabolically stabilized LPA analog, OTP, to induce LPA signaling (39, 40). Of note, OTP is preferentially recognized by LPA5 relative to other LPA receptors and displays a much lower EC50 for LPA5 compared with other LPA receptors (41).
Using this system, we monitored CD8 T-cell proliferation and expression of activation markers after antigen-specific TCR stimulation in vitro in the absence or presence of LPA signaling achieved with OTP. Proliferation was monitored by CFSE dilution and showed that in the presence of OTP, CD8 T cells were acutely inhibited from proliferating in response to G4 peptide stimulation (Fig. 2A). Specifically, by day 3, control cells had typically started to divide with the majority having undergone at least two divisions by day 7, whereas the majority of OTP-treated cells remained undivided in the same period. OTP inhibition of CD8 T-cell activation was further evidenced by significantly reduced expression of both the CD25 and CD44 activation antigens after TCR stimulation (Fig. 2B). Likely as a consequence of inhibiting activation and proliferation, OTP treatment also resulted in a reduced accumulation of CD8+ T cells (Fig. 2C) without any significant effect on viability as compared with unstimulated controls (Fig. 2D). Importantly, OTP treatment did not affect the ability of antigen-presenting cells to present peptide to OT-I T cells as OTP-treated and -untreated antigen-presenting cells were equally able to induce CD25 upregulation (Supplementary Fig. S3). Similar to our initial experiment with anti-CD3 stimulation, we found that the ability of LPA signaling to dampen CD8 T-cell proliferation and activation was less effective after the TCR was stimulated with the higher affinity SIINFEKL peptide (Fig. 2E). In addition, OTP treatment did not seem to significantly influence SIINFEKL-stimulated TNF-α or IFN-γ cytokine production by in vitro–generated effector CD8+ T cells (Fig. 2F). Together, these findings show that LPA signaling potently suppresses TCR signaling, cell activation, and proliferation when T cells are stimulated by relatively weak-affinity antigens.
Increased LPA signaling dampens activation and proliferation in vivo
Our in vitro data show that LPA signaling via LPA5 on CD8 T cells inhibits TCR signaling, activation, and proliferation. We next addressed whether this LPA regulatory pathway operated similarly in vivo. To accomplish this, we transferred purified, CFSE-labeled OT-I CD8+ T cells into C57BL/6 recipients followed by antigen-specific stimulation achieved by the subsequent transfer of BMDC previously pulsed with TCR-specific peptide (Fig. 3A). Under these conditions, BMDC pulsed with the relatively weak-affinity G4 peptide were unable to stimulate OT-I CD8+ T cells as determined by the absence of increased activation antigen expression or proliferation (data not shown). In contrast, when SIINFEKL peptide-pulsed BMDC (SIINFEKL-BMDC) were transferred, OT-I CD8 T-cell proliferation was observed clearly 3 days later in the dLN (axillary, brachial, and cervical) relative to the site of BMDC transfer (Fig. 3B).
To promote LPA signaling in vivo, we treated mice with OTP subcutaneously, which results in detectable levels in the blood after 1 hour with an approximate half-life of 5.5 hours (G. Tigyi; unpublished data). Mice were treated with 5 mg/kg OTP every 8 hours, with the first dose preceding SIINFEKL-BMDC transfer by 1 hour (Fig. 3A). Three days after antigen-specific stimulation, the OT-I CD8 T cells recovered from the dLN of OTP-treated animals had proliferated considerably less compared with those from the vehicle-treated animals (Fig. 3B). Furthermore, the number of OT-I CD8+ T cells recovered in the dLN of OTP-treated mice was significantly reduced relative to those recovered from the vehicle-treated animals and consistent with an inhibition of proliferation (Fig. 3C). In addition, antigen-specific T-cell activation was diminished, as evidenced by the reduced expression of CD25 by the transferred OT-I CD8 T cells in OTP-treated mice compared with those of the vehicle-treated mice (Fig. 3D). Taken together, these findings show that increased LPA signaling inhibits antigen-specific CD8 T-cell proliferation and activation both in vitro and in vivo.
LPA5-deficient CD8 T cells show enhanced proliferation in response to antigen-specific stimulation in vivo
To directly test whether LPA5 signaling suppressed CD8 T cells in vivo, we measured antigen-driven proliferation of LPA5-deficient and wild-type OT-I CD8 T cells after adoptive transfer. LPA5−/− mice are phenotypically unremarkable compared with wild-type littermates and display comparable numbers and frequencies of splenic CD8+ and CD4+ T-cell populations (Supplementary Fig. S4), similar to that reported for an independently generated LPA5-deficient mouse strain (42). Furthermore, in vitro LPA5-deficient CD8 T cells displayed similar viability as wild-type cells (data not shown). To address whether LPA inhibited antigen-specific CD8 T-cell responses under normal physiologic conditions, LPA5+/+ or LPA5−/− OT-I CD8+ T cells were isolated, CFSE-loaded, transferred into recipients, which were subsequently immunized with SIINFEKL-pulsed BMDC similar to those shown in Fig. 3A except neither group was treated with OTP. The results from these experiments show that 3 days after antigen-specific stimulation, a considerable proportion of wild-type OT-I CD8 T cells had undergone cell division. In contrast, similar stimulation of LPA5-deficient OT-I CD8 T cells resulted in an increased percentage of LPA5-deficient OT-I CD8 T cells that had undergone cell division and fewer transferred cells that remained CFSEhigh (Fig. 4A). Consistent with an increased frequency of LPA5−/− OT-I CD8 T cells having proliferated, higher numbers of mutant OT-I CD8 T cells were recovered from the dLN compared with wild-type cells (Fig. 4B). Together, these data reveal that in the absence of LPA5-mediated suppression, antigen-specific stimulation of OT-I CD8 T cells leads to a higher frequency of cells that proliferate and accumulate in the dLN.
Transfer of LPA5-deficient tumor-specific CD8 T cells controls tumor progression
Our data thus far show that LPA signaling via LPA5 expressed by CD8 T cells suppresses T-cell activation and proliferation. A number of different tumor cell types have been reported to produce elevated levels of LPA that promote survival, growth, and tumorigenesis (26, 43). Thus, we next compared the ability of LPA5-deficient and -sufficient tumor-specific CD8 T cells to control progression of an established tumor.
To address this question, we used the B16 melanoma cell line, B16.cOVA, which stably expresses chicken ovalbumin (with the SIINFEKL peptide) and used OT-I T cells as tumor-specific CD8 T cells as previously reported (44). B16.cOVA cells were subcutaneously implanted in the rear leg of wild-type C57BL/6 mice, and 5 days later either naïve LPA5−/− or wild-type OT-I CD8+ T cells were transferred into these recipients and the tumor diameter measured every 2 days thereafter (Fig. 5A). The results from these experiments showed that tumors grew similarly in the presence or absence of wild-type OT-I CD8 T cell transfer (Fig. 5B and E), consistent with previous reports showing tumor progression in the presence of wild-type tumor-specific CD8 T cells (45, 46). In contrast, tumor growth was clearly abated in mice 6 and 8 days after receiving LPA5-deficient OT-I CD8+ T cells (Fig. 5B and E). Compared with the wild-type tumor-specific CD8 T cells, LPA5-deficient tumor-specific CD8 T cells were also found typically at higher numbers within the tumor compared with wild-type (Fig. 5C). However, compared with the wild-type CD8 T cells, these LPA5-deficient tumor-infiltrating CD8 T cells seemed to express similar levels of IFN-γ, TNF-α, the inhibitory PD-1 receptor, and the CCR7 chemokine receptor 5 days after transfer (Fig. 5D and data not shown). Averaging tumor size across a treatment group showed that while the tumors were similar in size at the time of T-cell transfer, a significant reduction in tumor size was observed at 6 and 8 days posttransfer of LPA5−/− tumor-specific CD8 T cells compared with the tumor size in mice transferred with wild-type CD8 T cells (Fig. 5B). Further evidence that tumor-specific LPA5−/− CD8 T cells were better able to control tumor progression as compared with wild-type CD8 T cells was provided by the significantly smaller tumors that were harvested 8 days after T-cell transfer (Fig. 5E and F). These data show that tumor growth is inhibited to a greater degree when LPA5 signaling is prevented in tumor-specific CD8 T cells.
CD8 T cells are specialized cells of the adaptive immune system with the ability to recognize and eliminate nascent tumors. However, tumors are notorious for promoting an immunosuppressive environment through different mechanisms that thwart this adaptive immune response. Notably, the identification and targeting of some of these immunosuppressive mechanisms has led to relative success in the immunotherapeutic treatment of melanoma (3–5). Our data reveal a previously uncharacterized ability of the LPA lysophospholipid to suppress CD8 TCR signaling and in vivo activation, proliferation, and tumor control. As aberrant production of LPA is a common feature of diverse cancer cell types, these data further suggest that tumor cells can exploit a naturally occurring lipid to not only promote tumorigenesis but also to create an immunosuppressive environment.
Precedence for lysophospholipid regulation of adaptive immunity has been established previously in studies of S1P, which plays an important role in directing lymphocyte trafficking, localization, and development (47, 48). Similar to S1P, LPA is recognized by multiple GPCR that are differentially expressed by lymphocytes; in previous reports using primary human CD4 T cells and T-cell lines, LPA has been shown to regulate intracellular calcium mobilization (49) and cytokine production (16, 50). However, these in vitro studies did not identify the precise T-cell signaling pathway regulated by LPA, or which LPA receptor mediated these activities. In this report, we show that LPA engagement of LPA5 suppresses CD8+ T-cell receptor signaling, activation, and tumor immunity.
CD8 T cells express several LPA receptors and we have identified LPA5 expression to be required for not only negative regulation of TCR-induced calcium mobilization but also for inhibiting in vivo antigen-mediated proliferation. TCR-mediated increase in cytosolic calcium is an early signaling event important for both proximal and distal CD8 T-cell activities (33). Indeed, the primary encounter between antigen-specific CD8 T cells and the specific antigen is known to program the subsequent proliferation and differentiation of antigen-specific cells in vitro and in vivo (51, 52). Analyses of TCR-mediated intracellular calcium mobilization unambiguously showed the ability of LPA to suppress TCR signaling at 1 μmol/L, the lowest concentration tested, which approximates the physiologic level of LPA in normal individuals (18). At higher concentrations of LPA (or OTP), antigen-mediated CD8 T-cell activation and proliferation in vitro were considerably suppressed. However, it is not yet clear whether the source of LPA that suppresses CD8 T-cell tumor immunity is derived from the tumor, the endogenous systemic levels, or from elsewhere, which is currently under investigation in our laboratory. It is important to note that the significance of the systemic levels of LPA in regard to LPA receptor signaling is not yet established. Current models suggest that biologically relevant LPA signaling results from localized and directed autocrine/paracrine LPA production and signaling (19, 53) where concentrations can reach considerably higher levels. In this model, secreted autotaxin has been proposed to associate with integrins on a cell surface where LPA is locally produced and engages nearby LPA receptors.
To assess the ability of LPA–LPA5 signaling to suppress antigen-specific CD8 T-cell responses in vitro and in vivo, we used an established model system that allowed us to either increase LPA signaling using an LPA analogue, OTP, or prevent LPA inhibitory signaling using LPA5-deficient CD8 T cells. Our in vitro data suggest that LPA suppression of antigen-specific CD8 T cells operates more efficiently for relatively weak-affinity peptide antigens (Fig. 2 and Supplementary Fig. S2). However, in our studies neither wild-type nor LPA5−/− OT-I CD8 T cells were stimulated in vivo with BMDC pulsed with the lower-affinity G4 peptide. Thus, whether LPA5 suppression operates particularly for relatively weak affinity antigens could not be confirmed in vivo, but this is an area of ongoing investigation. As endogenous tumor-specific CD8 T cells normally express TCR with relatively low affinity for tumor antigens (54–56), this would suggest that tumor-derived LPA may be particularly effective at suppressing CD8 T-cell tumor immunity.
Our results clearly show that LPA signaling via LPA5 inhibits CD8 T-cell TCR signaling, activation, and proliferation and that LPA5-deficient tumor-specific CD8 T cells are better than the wild-type cells at controlling the growth of established tumor. However, analysis of tumor-infiltrating CD8 T cells 5 days after adoptive transfer did not reveal any apparent difference between tumor-specific LPA5−/− and wild-type CD8 T cells with respect to the expression of IFN-γ, TNF-α, PD-1 inhibitory receptor, or the CCR7 chemokine receptor. Nevertheless, when stimulated with specific antigen in vivo, LPA5-deficient CD8 T cells consistently proliferated to a greater extent and accumulated to higher numbers relative to wild-type CD8 T cells (Figs. 4 and 5C). Thus, together these data support a model in which LPA5−/− tumor-specific CD8 T cells are more easily stimulated to proliferate in response to tumor antigen and the increased proliferation and accumulation equate, at least in part, to better tumor control. We note, however, that LPA has been shown to influence T-cell homing (57) and migration (58). Thus, it remains possible that LPA5−/− tumor-specific CD8 T cells also display altered migration and trafficking, or they could influence CD8 T-cell function by other indirect mechanism(s) and, which is the focus of ongoing experiments.
As enhanced LPA production is a feature of many malignant cell types, our findings further suggest that in addition to the role of this lipid in enhancing tumorigenesis (27–29), LPA production by these cells also represents an additional inhibitory mechanism within the tumor microenvironment that serves to impede tumor rejection by CD8 T cells. The recent success of immunotherapies that antagonize inhibitory receptor signaling by T cells suggests that identification and characterization of additional signaling pathways used by tumors to suppress CD8 T-cell tumor immunity will only improve cancer immunotherapy (6, 7). Data presented here show that LPA5 functions as an additional inhibitory receptor on CD8 T cells. Given the association of LPA with multiple cancers (26, 43, 59), blockade of LPA5 signaling may be a promising additional strategy to promote host CD8 T-cell tumor immunity.
Disclosure of Potential Conflicts of Interest
T. Oravecz has ownership interest (including patents) in Lexicon Pharmaceuticals. G. Tigyi has ownership interest (including patents) in RxBio Inc. and is a consultant/advisory board member of the same. No potential conflicts of interest were disclosed by the other authors.
Conception and design: S.K. Oda, G. Tigyi, R. Pelanda, R.M. Torres
Development of methodology: S.K. Oda, P. Strauch, G. Tigyi, R.M. Torres
Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): S.K. Oda, P. Strauch, Y. Fujiwara, A. Al-Shami, T. Oravecz, R.M. Torres
Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): S.K. Oda, R.M. Torres
Writing, review, and/or revision of the manuscript: S.K. Oda, A. Al-Shami, T. Oravecz, G. Tigyi, R. Pelanda, R.M. Torres
Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): S.K. Oda, P. Strauch, R.M. Torres
Study supervision: R. Pelanda, R.M. Torres
This work was supported by the NIH (AI052157, to R.M. Torres and AI08405, to G. Tigyi), Cancer League of Colorado (to R.M. Torres), and Cancer Research Institute Special Emphasis Program in Tumor Immunology Award (to S.K. Oda).
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
The authors thank Dr. Jerold Chun (Scripps Research Institute) for LPA2-deficient mice and Dr. Dennis Voelker and his laboratory (NJH) for help with lipid handling. The authors also thank the R&R Laboratory for constructive criticisms and suggestions; Carrie Happoldt for excellent technical assistance with animal experiments; Matthew A. Burchill, Jason Z. Oh, and Jonathan D. Buhrman for helpful discussions; and the BRC staff and the staff at the NJH Cytometry Core for their assistance.
Note: Supplementary data for this article are available at Cancer Immunology Research Online (http://cancerimmunolres.aacrjournals.org/).
- Received July 22, 2013.
- Accepted July 25, 2013.
- ©2013 American Association for Cancer Research.