Advertisement
Research Articles

Lysophosphatidic Acid Inhibits CD8 T-cell Activation and Control of Tumor Progression

Shannon K. Oda, Pamela Strauch, Yuko Fujiwara, Amin Al-Shami, Tamas Oravecz, Gabor Tigyi, Roberta Pelanda and Raul M. Torres
DOI: 10.1158/2326-6066.CIR-13-0043-T Published October 2013

Abstract

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.

Introduction

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

Mice

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.

Calcium mobilization

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.

qPCR

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.

Cytometry

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).

Lipid preparation

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.

Statistical analysis

All statistical analysis was conducted with GraphPad Prism software (version 5.0) using a two-tailed unpaired Student t test.

Results

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).

Figure 1.
Figure 1.

LPA signals via LPA5 to inhibit CD8+ T-cell TCR–mediated intracellular calcium release. A, intracellular calcium concentration was assessed in purified CD8+ T cells stimulated with 10 μg/mL of anti-CD3 in the absence (black line) or presence of the indicated titrated concentrations of LPA. B, expression of Lpar1-6 by isolated CD8+ T cells as measured by qPCR. C, LPA2−/− or LPA5−/− CD8+ T cells were isolated, and intracellular calcium mobilization was assessed in response to anti-CD3 stimulation in the presence (bold line) or absence (thin line) of 20 μmol/L LPA.

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.

Figure 2.
Figure 2.

LPA signaling inhibits in vitro TCR-induced activation and proliferation. A, CD8+ T cells were stimulated with specific antigen in the absence (top row) or presence (middle row) of OTP and analyzed for expression of the CD25 activation antigen and CFSE fluorescence at the indicated times. Overlay of CFSE histograms by day is shown in bottom row in the absence (thin line) or presence of OTP (bold line) or when left unstimulated (Unstim.; gray filled). Representative of three independent experiments. B, CD25 and CD44 expression on CD8+ T cells is shown in histograms after 3 or 4 days, respectively, of in vitro culture in the absence of stimulation (gray filled) or after G4 peptide stimulation alone (thin line), or in the presence of OTP (bold line). Histograms are representative of three independent experiments, and normalized MFI values are summarized in bar chart (mean + SEM; **, P < 0.01; ***, P < 0.0001). C, enumeration of in vitro–cultured peptide-specific OT-I CD8+ T cells at the indicated times in the absence of peptide (gray circles) or after peptide stimulation alone (open circles) or in the presence of 20 μmol/L OTP (black circles). D, cell viability was determined by 7-AAD staining at the peak of CD25 expression, and data are representative of three independent experiments (mean + SEM); ns, not statistically significant. E, splenocytes from OT-I transgenic mice were labeled with CFSE, stimulated with SIINFEKL peptide, and cultured for 3 days before analyses of CFSE fluorescence. F, effector OT-I CD8+ T cells were left unstimulated (left) or restimulated with SIINFEKL in the absence (center) or presence (right) of OTP before analysis of TNF-α and IFN-γ expression. Data are representative of two independent experiments.

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).

Figure 3.
Figure 3.

LPA signaling inhibits TCR-mediated CD8+ T cell activation in vivo. A, schematic of adoptive transfer of OT-I CD8+ T cells and in vivo OTP treatment. B, histograms of CFSE dilution in vehicle-treated control (thin line), OTP-treated (bold line), and unstimulated control (gray filled) OT-I CD8+ T cells isolated from dLN. C, number of OT-I CD8+ T cells in dLN. D, representative histograms of CD25 expression by OT-I CD8+ T cells after vehicle-treatment (thin line), OTP-treatment (bold line), or unstimulated (gray shaded). Right, expression of CD25 as MFI by OT-I CD8+ T cells in the dLN from individual mice after vehicle treatment (closed circles) or OTP treatment (open circles). Data in B, C, and D are representative of two independent experiments (n = 4 mice/group; mean + SEM; **, P < 0.01).

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.

Figure 4.
Figure 4.

LPA5-deficient CD8+ T cells stimulated by peptide in vivo exhibit enhanced proliferation and accumulation. A and B, purified LPA5+/+ or LPA5−/− OT-I CD8+ T cells were CFSE labeled and transferred to C57BL/6 recipients, similar to Fig. 3A. Recipients were immunized 1 day later with SIINFEKL-pulsed BMDCs. On day 3 after immunization, mice were sacrificed and draining lymph nodes harvested for analysis. A, proliferation of LPA5+/+ (thin line), LPA5−/− (bold line), or unimmunized (filled gray) OT-I CD8+ T cells as indicated by CFSE dilution. Data are representative of two independent experiments. B, number of OT-I CD8+ T cells harvested from the dLN of LPA5+/+ (closed circles; n = 3–4) and LPA5−/− (open circles; n = 3–4) OT-I CD8+ T-cell recipients in the absence of stimulation or 3 days after peptide stimulation (**, P < 0.01). Data are representative of two independent experiments.

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.

Figure 5.
Figure 5.

Enhanced control of tumor progression by LPA5-deficient CD8+ T cells. A, schematic of tumor establishment and subsequent OT-I CD8+ T-cell adoptive transfer. C57BL/6 recipients were implanted subcutaneously in the leg with 105 B16.cOVA cells. Five days after implantation, 0.5 × 106 LPA5+/+ or LPA5−/− OT-I CD8+ T cells were transferred intravenously and tumor progression evaluated at the specified time points. B, average combined tumor burden at the indicated days in the absence of T-cell transfer (gray circles; n = 9) or after adoptive transfer of LPA5+/+ OT-I (black; n = 10), LPA5−/− OT-I (white; n = 11), or untreated controls (gray; n = 9). Data are cumulative of three independent experiments (mean ± SEM; *, P < 0.05; **, P < 0.01). C, number of tumor-specific CD8+ T cells per gram of tumor at day 5 after adoptive transfer of LPA5+/+ (black; n = 11) or LPA5−/− (white; n = 11) OT-I CD8+ T cells. Data are cumulative of three independent experiments (mean ± SEM; P = 0.08). D, tumors were harvested on day 5 after T-cell transfer and LPA5+/+ (black line) and LPA5−/− (dotted line) OT-I CD8 T cells evaluated for IFN-γ and TNF-α expression and compared with tumor-free wild-type lymph node OT-I T cells (gray histogram). E, tumor mass 8 days after adoptive transfer of LPA5+/+ OT-I (black; n = 10), LPA5−/− OT-I (white; n = 11), or untreated controls (gray; n = 9). Data are cumulative of three independent experiments (mean ± SEM; **, P < 0.01). F, representative tumors in situ and ex vivo from C57BL/6 mice in the absence of T-cell transfer or 8 days after adoptive transfer of LPA5+/+ or LPA5−/− tumor-specific CD8+ T cells.

Discussion

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.

Authors' Contributions

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

Grant Support

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.

Acknowledgments

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.

Footnotes

  • 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.

References

  1. 1.↵
    1. Dunn GP,
    2. Old LJ,
    3. Schreiber RD
    . The immunobiology of cancer immunosurveillance and immunoediting. Immunity 2004;21:137–48.
    OpenUrlCrossRefPubMed
  2. 2.↵
    1. Schreiber RD,
    2. Old LJ,
    3. Smyth MJ
    . Cancer immunoediting: integrating immunity's roles in cancer suppression and promotion. Science 2011;331:1565–70.
    OpenUrlAbstract/FREE Full Text
  3. 3.↵
    1. Hodi FS,
    2. O'Day SJ,
    3. McDermott DF,
    4. Weber RW,
    5. Sosman JA,
    6. Haanen JB,
    7. et al.
    Improved survival with ipilimumab in patients with metastatic melanoma. N Engl J Med 2010;363:711–23.
    OpenUrlCrossRefPubMed
  4. 4.↵
    1. Hamid O,
    2. Robert C,
    3. Daud A,
    4. Hodi FS,
    5. Hwu WJ,
    6. Kefford R,
    7. et al.
    Safety and tumor responses with lambrolizumab (Anti-PD-1) in melanoma. N Engl J Med 2013;369:134–44.
    OpenUrlCrossRefPubMed
  5. 5.↵
    1. Wolchok JD,
    2. Kluger H,
    3. Callahan MK,
    4. Postow MA,
    5. Rizvi NA,
    6. Lesokhin AM,
    7. et al.
    Nivolumab plus ipilimumab in advanced melanoma. N Engl J Med 2013;369:122–33.
    OpenUrlCrossRefPubMed
  6. 6.↵
    1. Pardoll DM
    . The blockade of immune checkpoints in cancer immunotherapy. Nat Rev Cancer 2012;12:252–64.
    OpenUrlCrossRefPubMed
  7. 7.↵
    1. Stewart TJ,
    2. Smyth MJ
    . Improving cancer immunotherapy by targeting tumor-induced immune suppression. Cancer Metastasis Rev 2011;30:125–40.
    OpenUrlCrossRefPubMed
  8. 8.↵
    1. Zehn D,
    2. King C,
    3. Bevan MJ,
    4. Palmer E
    . TCR signaling requirements for activating T cells and for generating memory. Cell Mol Life Sci 2012;69:1565–75.
    OpenUrlCrossRefPubMed
  9. 9.↵
    1. Smith-Garvin JE,
    2. Koretzky GA,
    3. Jordan MS
    . T cell activation. Ann Rev Immunol 2009;27:591–619.
    OpenUrlCrossRefPubMed
  10. 10.↵
    1. Peggs KS,
    2. Quezada SA,
    3. Allison JP
    . Cell intrinsic mechanisms of T-cell inhibition and application to cancer therapy. Immunol Rev 2008;224:141–65.
    OpenUrlCrossRefPubMed
  11. 11.↵
    1. Leach DR,
    2. Krummel MF,
    3. Allison JP
    . Enhancement of antitumor immunity by CTLA-4 blockade. Science 1996;271:1734–6.
    OpenUrlAbstract
  12. 12.↵
    1. Tigyi G
    . Aiming drug discovery at lysophosphatidic acid targets. Br J Pharmacol 2010;161:241–70.
    OpenUrlCrossRefPubMed
  13. 13.↵
    1. Blaho VA,
    2. Hla T
    . Regulation of mammalian physiology, development, and disease by the sphingosine 1-phosphate and lysophosphatidic acid receptors. Chem Rev 2011;111:6299–320.
    OpenUrlCrossRefPubMed
  14. 14.↵
    1. Chun J,
    2. Hla T,
    3. Lynch KR,
    4. Spiegel S,
    5. Moolenaar WH
    . International Union of Basic and Clinical Pharmacology. LXXVIII. Lysophospholipid receptor nomenclature. Pharmacol Rev 2010;62:579–87.
    OpenUrlAbstract/FREE Full Text
  15. 15.↵
    1. Kotarsky K,
    2. Boketoft A,
    3. Bristulf J,
    4. Nilsson NE,
    5. Norberg A,
    6. Hansson S,
    7. et al.
    Lysophosphatidic acid binds to and activates GPR92, a G protein–coupled receptor highly expressed in gastrointestinal lymphocytes. J Pharmacol Exp Ther 2006;318:619–28.
    OpenUrlAbstract/FREE Full Text
  16. 16.↵
    1. Goetzl EJ,
    2. Kong Y,
    3. Voice JK
    . Cutting edge: differential constitutive expression of functional receptors for lysophosphatidic acid by human blood lymphocytes. J Immunol 2000;164:4996–9.
    OpenUrlAbstract/FREE Full Text
  17. 17.↵
    1. Choi JW,
    2. Herr DR,
    3. Noguchi K,
    4. Yung YC,
    5. Lee CW,
    6. Mutoh T,
    7. et al.
    LPA receptors: subtypes and biological actions. Annu Rev Pharmacol Toxicol 2010;50:157–86.
    OpenUrlCrossRefPubMed
  18. 18.↵
    1. Smyth SS,
    2. Cheng HY,
    3. Miriyala S,
    4. Panchatcharam M,
    5. Morris AJ
    . Roles of lysophosphatidic acid in cardiovascular physiology and disease. Biochim Biophys Acta 2008;1781:563–70.
    OpenUrlPubMed
  19. 19.↵
    1. Moolenaar WH,
    2. Perrakis A
    . Insights into autotaxin: how to produce and present a lipid mediator. Nat Rev Mol Cell Biol 2011;12:674–9.
    OpenUrlCrossRefPubMed
  20. 20.↵
    1. Stracke ML,
    2. Krutzsch HC,
    3. Unsworth EJ,
    4. Arestad A,
    5. Cioce V,
    6. Schiffmann E,
    7. et al.
    Identification, purification, and partial sequence analysis of autotaxin, a novel motility-stimulating protein. J Biol Chem 1992;267:2524–9.
    OpenUrlAbstract/FREE Full Text
  21. 21.↵
    1. Houben AJ,
    2. Moolenaar WH
    . Autotaxin and LPA receptor signaling in cancer. Cancer Metastasis Rev 2011;30:557–65.
    OpenUrlCrossRefPubMed
  22. 22.↵
    1. Samadi N,
    2. Bekele R,
    3. Capatos D,
    4. Venkatraman G,
    5. Sariahmetoglu M,
    6. Brindley DN
    . Regulation of lysophosphatidate signaling by autotaxin and lipid phosphate phosphatases with respect to tumor progression, angiogenesis, metastasis and chemo-resistance. Biochimie 2011;93:61–70.
    OpenUrlCrossRefPubMed
  23. 23.↵
    1. Gotoh M,
    2. Fujiwara Y,
    3. Yue J,
    4. Liu J,
    5. Lee S,
    6. Fells J,
    7. et al.
    Controlling cancer through the autotaxin-lysophosphatidic acid receptor axis. Biochem Soc Trans 2012;40:31–6.
    OpenUrlAbstract/FREE Full Text
  24. 24.↵
    1. Westermann AM,
    2. Havik E,
    3. Postma FR,
    4. Beijnen JH,
    5. Dalesio O,
    6. Moolenaar WH,
    7. et al.
    Malignant effusions contain lysophosphatidic acid (LPA)-like activity. Ann Oncol 1998;9:437–42.
    OpenUrlAbstract/FREE Full Text
  25. 25.↵
    1. Xiao YJ,
    2. Schwartz B,
    3. Washington M,
    4. Kennedy A,
    5. Webster K,
    6. Belinson J,
    7. et al.
    Electrospray ionization mass spectrometry analysis of lysophospholipids in human ascitic fluids: comparison of the lysophospholipid contents in malignant vs nonmalignant ascitic fluids. Anal Biochem 2001;290:302–13.
    OpenUrlCrossRefPubMed
  26. 26.↵
    1. Xu Y,
    2. Shen Z,
    3. Wiper DW,
    4. Wu M,
    5. Morton RE,
    6. Elson P,
    7. et al.
    Lysophosphatidic acid as a potential biomarker for ovarian and other gynecologic cancers. JAMA 1998;280:719–23.
    OpenUrlCrossRefPubMed
  27. 27.↵
    1. Shida D,
    2. Kitayama J,
    3. Yamaguchi H,
    4. Okaji Y,
    5. Tsuno NH,
    6. Watanabe T,
    7. et al.
    Lysophosphatidic acid (LPA) enhances the metastatic potential of human colon carcinoma DLD1 cells through LPA1. Cancer Res 2003;63:1706–11.
    OpenUrlAbstract/FREE Full Text
  28. 28.↵
    1. Bian D,
    2. Su S,
    3. Mahanivong C,
    4. Cheng RK,
    5. Han Q,
    6. Pan ZK,
    7. et al.
    Lysophosphatidic acid stimulates ovarian cancer cell migration via a Ras–MEK kinase 1 pathway. Cancer Res 2004;64:4209–17.
    OpenUrlAbstract/FREE Full Text
  29. 29.↵
    1. Rivera-Lopez CM,
    2. Tucker AL,
    3. Lynch KR
    . Lysophosphatidic acid (LPA) and angiogenesis. Angiogenesis 2008;11:301–10.
    OpenUrlCrossRefPubMed
  30. 30.↵
    1. Vidot S,
    2. Witham J,
    3. Agarwal R,
    4. Greenhough S,
    5. Bamrah HS,
    6. Tigyi GJ,
    7. et al.
    Autotaxin delays apoptosis induced by carboplatin in ovarian cancer cells. Cell Signal 2010;22:926–35.
    OpenUrlCrossRefPubMed
  31. 31.↵
    1. Brindley DN,
    2. Lin FT,
    3. Tigyi GJ
    . Role of the autotaxin-lysophosphatidate axis in cancer resistance to chemotherapy and radiotherapy. Biochim Biophys Acta 2013;1831:74–85.
    OpenUrlCrossRefPubMed
  32. 32.↵
    1. Durgam GG,
    2. Virag T,
    3. Walker MD,
    4. Tsukahara R,
    5. Yasuda S,
    6. Liliom K,
    7. et al.
    Synthesis, structure–activity relationships, and biological evaluation of fatty alcohol phosphates as lysophosphatidic acid receptor ligands, activators of PPARgamma, and inhibitors of autotaxin. J Med Chem 2005;48:4919–30.
    OpenUrlCrossRefPubMed
  33. 33.↵
    1. Hogan PG,
    2. Lewis RS,
    3. Rao A
    . Molecular basis of calcium signaling in lymphocytes: STIM and ORAI. Annu Rev Immunol 2010;28:491–533.
    OpenUrlCrossRefPubMed
  34. 34.↵
    1. Jung TM,
    2. Gallatin WM,
    3. Weissman IL,
    4. Dailey MO
    . Down-regulation of homing receptors after T cell activation. J Immunol 1988;141:4110–7.
    OpenUrlAbstract
  35. 35.↵
    1. Hogquist KA,
    2. Jameson SC,
    3. Heath WR,
    4. Howard JL,
    5. Bevan MJ,
    6. Carbone FR
    . T cell receptor antagonist peptides induce positive selection. Cell 1994;76:17–27.
    OpenUrlCrossRefPubMed
  36. 36.↵
    1. Zehn D,
    2. Lee SY,
    3. Bevan MJ
    . Complete but curtailed T-cell response to very low-affinity antigen. Nature 2009;458:211–4.
    OpenUrlCrossRefPubMed
  37. 37.↵
    1. Rosette C,
    2. Werlen G,
    3. Daniels MA,
    4. Holman PO,
    5. Alam SM,
    6. Travers PJ,
    7. et al.
    The impact of duration versus extent of TCR occupancy on T cell activation: a revision of the kinetic proofreading model. Immunity 2001;15:59–70.
    OpenUrlCrossRefPubMed
  38. 38.↵
    1. Albers HM,
    2. Dong A,
    3. van Meeteren LA,
    4. Egan DA,
    5. Sunkara M,
    6. van Tilburg EW,
    7. et al.
    Boronic acid-based inhibitor of autotaxin reveals rapid turnover of LPA in the circulation. Proc Natl Acad Sci U S A 2010;107:7257–62.
    OpenUrlAbstract/FREE Full Text
  39. 39.↵
    1. Deng W,
    2. Shuyu E,
    3. Tsukahara R,
    4. Valentine WJ,
    5. Durgam G,
    6. Gududuru V,
    7. et al.
    The lysophosphatidic acid type 2 receptor is required for protection against radiation-induced intestinal injury. Gastroenterology 2007;132:1834–51.
    OpenUrlCrossRefPubMed
  40. 40.↵
    1. Kosanam H,
    2. Ma F,
    3. He H,
    4. Ramagiri S,
    5. Gududuru V,
    6. Tigyi GJ,
    7. et al.
    Development of an LC-MS/MS assay to determine plasma pharmacokinetics of the radioprotectant octadecenyl thiophosphate (OTP) in monkeys. J Chromatogr B Analyt Technol Biomed Life Sci 2010;878:2379–83.
    OpenUrlCrossRefPubMed
  41. 41.↵
    1. Kiss GN,
    2. Fells JI,
    3. Gupte R,
    4. Lee SC,
    5. Liu J,
    6. Nusser N,
    7. et al.
    Virtual screening for LPA2-specific agonists identifies a nonlipid compound with antiapoptotic actions. Mol Pharmacol 2012;82:1162–73.
    OpenUrlAbstract/FREE Full Text
  42. 42.↵
    1. Lin ME,
    2. Rivera RR,
    3. Chun J
    . Targeted deletion of LPA5 identifies novel roles for lysophosphatidic acid signaling in development of neuropathic pain. J Biol Chem 2012;287:17608–17.
    OpenUrlAbstract/FREE Full Text
  43. 43.↵
    1. Sasagawa T,
    2. Okita M,
    3. Murakami J,
    4. Kato T,
    5. Watanabe A
    . Abnormal serum lysophospholipids in multiple myeloma patients. Lipids 1999;34:17–21.
    OpenUrlCrossRefPubMed
  44. 44.↵
    1. Thompson ED,
    2. Enriquez HL,
    3. Fu YX,
    4. Engelhard VH
    . Tumor masses support naive T cell infiltration, activation, and differentiation into effectors. J Exp Med 2010;207:1791–804.
    OpenUrlAbstract/FREE Full Text
  45. 45.↵
    1. Nelson DJ,
    2. Mukherjee S,
    3. Bundell C,
    4. Fisher S,
    5. van Hagen D,
    6. Robinson B
    . Tumor progression despite efficient tumor antigen cross-presentation and effective “arming” of tumor antigen-specific CTL. J Immunol 2001;166:5557–66.
    OpenUrlAbstract/FREE Full Text
  46. 46.↵
    1. Blohm U,
    2. Roth E,
    3. Brommer K,
    4. Dumrese T,
    5. Rosenthal FM,
    6. Pircher H
    . Lack of effector cell function and altered tetramer binding of tumor-infiltrating lymphocytes. J Immunol 2002;169:5522–30.
    OpenUrlAbstract/FREE Full Text
  47. 47.↵
    1. Donovan EE,
    2. Pelanda R,
    3. Torres RM
    . S1P3 confers differential S1P-induced migration by autoreactive and non-autoreactive immature B cells and is required for normal B-cell development. Eur J Immunol 2010;40:688–98.
    OpenUrlCrossRefPubMed
  48. 48.↵
    1. Cyster JG,
    2. Schwab SR
    . Sphingosine-1-phosphate and lymphocyte egress from lymphoid organs. Annu Rev Immunol 2012;30:69–94.
    OpenUrlCrossRefPubMed
  49. 49.↵
    1. Wang L,
    2. Knudsen E,
    3. Jin Y,
    4. Gessani S,
    5. Maghazachi AA
    . Lysophospholipids and chemokines activate distinct signal transduction pathways in T helper 1 and T helper 2 cells. Cell Signal 2004;16:991–1000.
    OpenUrlCrossRefPubMed
  50. 50.↵
    1. Rubenfeld J,
    2. Guo J,
    3. Sookrung N,
    4. Chen R,
    5. Chaicumpa W,
    6. Casolaro V,
    7. et al.
    Lysophosphatidic acid enhances interleukin-13 gene expression and promoter activity in T cells. Am J Physiol Lung Cell Mol Physiol 2006;290:L66–74.
    OpenUrlAbstract/FREE Full Text
  51. 51.↵
    1. van Stipdonk MJ,
    2. Lemmens EE,
    3. Schoenberger SP
    . Naive CTLs require a single brief period of antigenic stimulation for clonal expansion and differentiation. Nat Immunol 2001;2:423–9.
    OpenUrlCrossRefPubMed
  52. 52.↵
    1. Kaech SM,
    2. Ahmed R
    . Memory CD8+ T cell differentiation: initial antigen encounter triggers a developmental program in naive cells. Nat Immunol 2001;2:415–22.
    OpenUrlCrossRefPubMed
  53. 53.↵
    1. Tabchy A,
    2. Tigyi G,
    3. Mills GB
    . Location, location, location: a crystal-clear view of autotaxin saturating LPA receptors. Nat Struct Mol Biol 2011;18:117–8.
    OpenUrlCrossRefPubMed
  54. 54.↵
    1. Slingluff CL Jr.,
    2. Hunt DF,
    3. Engelhard VH
    . Direct analysis of tumor-associated peptide antigens. Curr Opin Immunol 1994;6:733–40.
    OpenUrlCrossRefPubMed
  55. 55.↵
    1. Gervois N,
    2. Guilloux Y,
    3. Diez E,
    4. Jotereau F
    . Suboptimal activation of melanoma infiltrating lymphocytes (TIL) due to low avidity of TCR/MHC-tumor peptide interactions. J Exp Med 1996;183:2403–7.
    OpenUrlAbstract/FREE Full Text
  56. 56.↵
    1. Colella TA,
    2. Bullock TN,
    3. Russell LB,
    4. Mullins DW,
    5. Overwijk WW,
    6. Luckey CJ,
    7. et al.
    Self-tolerance to the murine homologue of a tyrosinase-derived melanoma antigen: implications for tumor immunotherapy. J Exp Med 2000;191:1221–32.
    OpenUrlAbstract/FREE Full Text
  57. 57.↵
    1. Zhang Y,
    2. Chen YC,
    3. Krummel MF,
    4. Rosen SD
    . Autotaxin through lysophosphatidic acid stimulates polarization, motility, and transendothelial migration of naive T cells. J Immunol 2012;189:3914–24.
    OpenUrlAbstract/FREE Full Text
  58. 58.↵
    1. Zheng Y,
    2. Kong Y,
    3. Goetzl EJ
    . Lysophosphatidic acid receptor-selective effects on Jurkat T cell migration through a Matrigel model basement membrane. J Immunol 2001;166:2317–22.
    OpenUrlAbstract/FREE Full Text
  59. 59.↵
    1. Mills GB,
    2. Moolenaar WH
    . The emerging role of lysophosphatidic acid in cancer. Nat Rev Cancer 2003;3:582–91.
    OpenUrlCrossRefPubMed
Back to top
Cancer Immunology Research: 1 (4)
October 2013
Volume 1, Issue 4

Sign up for alerts

View this article with LENS

Open full page PDF
Article Alerts
Email Article
Citation Tools
Lysophosphatidic Acid Inhibits CD8 T-cell Activation and Control of Tumor Progression
Shannon K. Oda, Pamela Strauch, Yuko Fujiwara, Amin Al-Shami, Tamas Oravecz, Gabor Tigyi, Roberta Pelanda and Raul M. Torres
Cancer Immunol Res October 1 2013 (1) (4) 245-255; DOI: 10.1158/2326-6066.CIR-13-0043-T
del.icio.us logo Digg logo Reddit logo Twitter logo CiteULike logo Facebook logo Google logo Mendeley logo

Jump to section

Advertisement