1. IntroductionDespite recent advances in diagnosing and treating gastric cancer, it remains the third most common cause of cancer-related death globally, especially in Asia, Europe, and the United States [1]. Distinct phenotypes of gastric cancer occur due to specific genetic alterations and epigenetic changes. Although different groups have reported the dysregulation of many signaling pathways, proteins, and multiple genes in gastric cancer, its mechanisms of carcinogenesis and heterogeneity remain unclear [2,3]. About 70% of gastric cancer patients show the dysregulation of the three major signaling pathways, including the Wnt β-catenin pathway, the nuclear factor κB (NFκB) pathway, and proliferation/stem cell pathways. Among these, dysregulated β-catenin pathways have been reported in 30–50% of gastric cancer patients and in different gastric cancer cell lines [4,5]. In addition, other types of human cancers are initiated by the mutation of other components of the β-catenin signaling pathway, such as β-catenin, adenomatous polyposis coli (APC), and Axin [6,7].Several studies have demonstrated that β-catenin signaling is activated by the mutation of these components and a variety of extracellular stimuli [7,8]. For example, Wnt proteins are well-known proteins that bind with the Frizzled receptors and activate the β-catenin pathway [9]. In addition, recent studies have revealed that β-catenin signaling is activated by GPCR signaling, and LPA acts through its G-protein-coupled receptor [8].Lysophosphatidic acid (LPA) is a multifunctional endogenous phospholipid that mediates the proliferation, differentiation, migration, regulation of cell–cell interaction [10], inhibition of cell death [11], neurite retraction [12], smooth muscle cell contraction [13], and transformation and progression in cancer [14]. LPA binds mainly with its six specific G-protein-coupled receptors (GPCR) to mediate the cellular responses. The aberrant expression of LPA receptors, particularly LPAR2 and LPAR3, has been reported in several cancers, suggesting a significant role for LPA in different cancer types through distinct signaling pathways such as AKT, MAPK, and ERK [15]. In this study, we show for the first time that LPA stimulates the proliferation, migration, and invasion of gastric cancer through the β-catenin signaling pathway and by increasing the energy metabolism, providing a novel insight into the molecular mechanism of gastric cancer. 2. Materials and Methods 2.1. Materials

LPA was purchased from Avanti Polar Lipids (Birmingham, AL, USA). Ki16425, an LPA antagonist against LPAR1, LPAR2, and LPAR3, was purchased from Echelon (Salt Lake City, UT, USA), and the LPAR2 antagonist (Cat # HY-18075) was purchased from MedChem Express (Monmouth Junction, NJ, USA) Rabbit monoclonal anti-β-catenin and rabbit polyclonal anti-LPA1# ab23698, LPA2#ab38322, LPA3#NBP1-84903, and LPP3 were purchased from Abcam (Cambridge, MA, USA). Antibodies against β-actin#3700 and GAPDH#97166 were purchased from Cell Signaling Technology (Beverly, MA, USA). Antibodies against c-Myc#13987T were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). Tissue lysates for the total protein were purchased from Novus Biologicals, LLC (Cat #NBP2-47080, NBP2 47081, and NBP2 47082). cDNA (Cat #C1234248, C1235248), RNA (Cat #R8235248-PP-10, R1235248-50, R1234253-10), and paraffin-embedded human tissue sections from the normal stomach and cancerous stomach (Cat# T8235248-PP) were purchased from Biochain (Newark, CA, USA). The LPA/Lysophosphatidic Acid (Competitive EIA) ELISA Kit-LS-F25111 was purchased from LSBio (Seattle, WA, USA).

2.2. Cell Culture

We purchased the AGS #CRL-1739 and NCI-N87 (ATCC Cat# CRL-5822, RRID: CVCL_1603) human gastric cancer cell lines from American Type Culture Collection (ATCC; Rockville, MD, USA). An ATCC-formulated F12 K medium was used for the AGS cell line and an RPMI-1640 medium was used for the NCI- N87 cell line. Both cell lines were supplemented with 10% fetal bovine serum (FBS) (Sigma, St. Louis, MO, USA), 100 U/mL penicillin, and 100 µg/mL streptomycin (Corning, NY, USA), maintained at a temperature of 37 °C and at 5% CO2 and sub-cultured by trypsinizing with trypsin-EDTA solution (Gibco, Gaithersburg, MD, USA).

2.3. LPA Treatment

The AGS and NCI-N87 cell lines were maintained throughout the study in the presence of 10% FBS F12K and 10% FBS RPMI medium, respectively. LPA (10 µM) was prepared with 1% (w/v) fatty acid-free BSA; before any treatment with LPA, the cells were starved in the presence of 0.1% dilapidated FBS F12 K or an RPMI medium for 24 h.

2.4. Cell Proliferation Assay

The cell proliferation assay was performed using the electric cell-substrate impedance sensing (ECIS) method, which measures the cell proliferation activity in real-time and by a hemocytometer. For the measurement using the hemocytometer, 0.5 × 105 cells were seeded in 6-well plates containing a complete growth medium and allowed to grow for 24 h. The cells were then treated with 10 µM of LPA after 24 h of starvation as described previously. The total number of cells in each well was counted every 24 h for 3 days after an LPA treatment. For the measurement using the ECIS proliferation assay, 15,000 cells were suspended in a complete growth medium were cultured on the surface of gold-film electrode-coated 8W20idf PET 8-well plates, which have ten small electrodes connected in parallel in each well. Once the cells were attached to the bottom, we treated the cells with the control (no LPA), 10 µM of LPA, an LPA + LPAR2 antagonist, or LPA+ Ki16425. The impedance was measured every 10 s at a frequency of 4000 for 60 h.

2.5. ECIS Wound-Healing Assay (ECIS)

An ECIS wound healing assay was performed using ECIS-Applied Biosystem technology. For the ECIS wound healing assay, 50,000 cells were seeded into the ECIS well and allowed to grow until a confluent monolayer formed and the cells entered the stationary phase. The cells were starved for 24 h after entering the stationary phase. After 24 h of starvation, electrical wounds were made in the ECIS plate by elevating the voltage pulse of a 40 kHz frequency at 3.5-V amplitude for 30 s. This sudden elevation in the voltage causes cell death and detachment from the active electrode. The medium was then aspirated and treated with 10 µM of LPA and the wound healing was assessed using continuous statistical analysis. The effect of Ki16425 and LPAR2 antagonists on the migration activity was measured using cells pretreated with 5 µM of LPAR2 antagonist or 10 µM of Ki16425 for 1 h and then exposed to 10 µM of the LPA +LPAR2 receptor antagonist or Ki16425.

2.6. Luciferase Reporter Assay

We transfected the AGS or NCI-N87 cells with the TOP flash luciferase reporter gene plasmid using the X-treme gene transfection reagent #0636624001(Rockville, MD, USA) according to the manufacturer’s instructions. We changed the medium of the transfected cells to a starvation medium after 24 h of transfection and maintained the cells under starvation conditions for 8 h. The serum-starved cells were treated with or without 10 µM of LPA for 24 h. For the treatment with inhibitors, serum-starved cells were pretreated with 10 µM of Ki16425 or 5 µM of LPAR2 antagonist for 1 h, followed by a co-treatment with 10 µM of LPA along with the respective inhibitors. Luminescence was measured in a single tube luminometer using the Promega luciferase assay system (E1500; Madison, WI, USA) and was normalized by the total protein concentration.

2.7. Mitochondrial Bioenergetics

We analyzed the OCR (oxygen consumption rate) and ECAR (extracellular acidification rate) using a Seahorse XF-24 extracellular flux analyzer (Seahorse Biosciences, Chicopee, MA, USA). Gastric cancer cells were grown in a Seahorse plate until they became fully confluent and treated with the control (no LPA), LPA (10 µM), or LPA plus the inhibitors. The bioenergetic activity of cells with or without an LPA treatment was monitored as free protons in real-time using the Seahorse XF-24 analyzer to measure the oxygen concentration. We quantified the OCR and ECAR values in pmol/min/μg and mpH/min/μg, respectively, and normalized the values to the total protein concentration. The initial basal value of the OCR was interrupted by the addition of oligomycin (Complex V inhibitor), giving values for the ATP-linked OCR. The addition of FCCP (an uncoupler) and rotenone + antimycin (Complex I and Complex III inhibitor) allowed us to determine the maximal OCR capacity and spare OCR capacity, respectively. We used a glucose-free medium for the ECAR analysis. Following the sequential addition of glucose (25 mM), oligomycin (1 μg/mL), and deoxyglucose (25 mM), we measured the rate of glycolysis and glycolytic reserve in the gastric cancer cells.

2.8. Data Analysis from the Clinical CohortWe used a gastric cancer TCGA data set obtained from the Cancer Genome Atlas (TCGA) through the UALCAN (UALCAN, RRID: SCR_015827; http://ualcan.path.uab.edu/cgi-bin/ualcan-res.pl; accessed on 1 November 2018) webserver to analyze the mRNA expression level of different LPA receptors and β-catenin mRNA levels [16]. 2.9. Statistical Analysis

Unless otherwise stated, the data are reported as means ± (SD). We repeated our in vitro experiments a minimum of three times. We used a one-way ANOVA with a Bonferroni correction for a significant correlation among the different groups. In addition, an unpaired Student’s t-test was used to identify the significant differences between the two groups. A statistical analysis was performed using prism 8.0 (GraphPad Prism, version 8.4.2, San Diego, CA, USA, RRID: SCR_002798).

4. DiscussionIn this report, we have demonstrated for the first time that LPA mediates the initiation and progression of gastric cancer through the LPAR2-β-catenin axis and by altering the energy metabolism. Existing studies in the literature suggest that a high LPA level plays a vital role in various kinds of tumor progression, such as ovarian carcinoma, thyroid carcinoma, pancreatic carcinoma, and colon cancer [24,25,26]. In addition, it has been shown that LPA mediates its downstream signaling pathway through GPCR and modulates various oncogenic signaling [27]. In agreement with this, the results from our clinical cohort studies revealed the high expression of LPAR2 in all stages of gastric cancer tissues compared to the expression in normal gastric tissues. These results suggest a possible role for LPA in the initiation and progression of gastric cancer through the LPAR2 receptor [17]. Data from another study have shown that LPAR-1, LPAR-2, and LPAR-3 are present in gastric cancer cell lines; among them, LPAR2 was expressed aberrantly in the AGS cell line [28]. However, the LPA-LPAR2-mediated functional effect and its downstream molecular signaling mechanism in gastric cancer remain to be elucidated. In this study, we demonstrated that LPA increased the proliferation and subsequent progression of the gastric cancer cells, which get abrogated into the functional level by both LPA receptor antagonists (Ki16425 and the LPAR2 receptor antagonist) and LPAR2 knockdown. Ki16425 is an antagonist of LPA1 and LPA3 with a moderate activity against LPA2 and the LPAR2 antagonist is specific to LPAR2. Since we observed a similar functional effect in both the LPA receptor antagonists and our cell line has a high expression of LPAR2, we speculated that LPA mediates its action through the LPAR2 receptor, which was confirmed by the LPAR2 knockdown.One of the most common causes of gastric carcinogenesis is the dysregulation of the β-catenin signaling pathway. β-Catenin is a multifunctional protein that maintains cell–cell adhesion by binding with E-cadherin and acts as a transcriptional regulator of β-catenin signaling. When the pathway is inactive, β-catenin binds with the destruction complex composed of Axin, APC, GSK-3β, and casein kinase 1α (CK1α). The phosphorylation of the destruction complex by the kinases leads to the degradation of cytoplasmic β-catenin, resulting in reduced β-catenin levels. In contrast, when the signaling pathway is active, it prevents the phosphorylation of β-catenin, leading to the subsequent stabilization of cytoplasmic β-catenin. A larger amount of cytoplasmic β-catenin then translocates into the nucleus and acts as a transcriptional stimulator of the target genes, such as Cyclin D1, c-MYC, and MMP2, by interacting with the TCF/LEF complex [29,30,31]. According to our findings, LPA induces the initiation and progression of gastric cancer by activating the β-catenin signaling pathway. On the other hand, the LPA-induced β-catenin activity was abrogated with LPA receptor antagonists. Furthermore, an LPA-induced β-catenin activation was confirmed by the knockdown of LPAR2 using LPAR2-shRNAs, which decreased the LPA-induced β-catenin activity.Since our findings revealed a functional role for LPA in gastric cancer through the activation of β-catenin and its downstream target genes, LPA may provide the oncogenic stimulation that increases the extent of a β-catenin activation and promotes the initiation and progression of gastric cancer. β-catenin cytoplasmic stabilization is due to the disassembly or phosphorylation of any of the β-catenin pathway components, such as APC, β-catenin, Axin, or GSK-3β. It has been shown that the mutation of exon 3 of β-catenin is the most common oncogenic step in human carcinogenesis, including gastric cancer [4]. However, Sekine et al. and Chan et al. showed that when only this deletion mutation occurs, it is not enough to induce carcinogenesis in the HCT116 colon cancer cell line [32,33]. LPA has been reported to phosphorylate GSK-3β in HEK 293 cells [34], and other groups have shown that the phosphorylation of only GSK-3β cannot sufficiently activate the β-catenin signaling pathway [35]. The exogenous overexpression of GSK-3β could not reduce the β-catenin-induced transcriptional activity in β-catenin overexpressing cells, whereas the β-catenin transcriptional activity was reduced after the addition of Axin or APC [36]. In this study, we have shown for the first time that LPA activates the β-catenin signaling pathway by increasing the phosphorylation of GSK-3β and by decreasing the mRNA level of Axin 2, which acts as a negative regulator of the β-catenin pathway (Supplementary Figure S4A,B). As a result, the cytoplasmic β-catenin destruction complex cannot be formed and β-catenin is stabilized. In 2004, Yang et al. showed that LPAR2 and LPAR3 bind to the Gαq and that LPA increased the phosphorylation of GSK-3β and the subsequent activation of the β-catenin signaling pathway in the HCT116 cell line through PKC3β1 [37]. Therefore, there is a possibility that LPAR2 increases the phosphorylation of GSK-3β by activating the protein kinase C, which means that further studies are required to elucidate the mechanism responsible for the LPA-LPAR2-induced phosphorylation of GSK-3β and its downstream β-catenin signaling pathway in gastric cancer.Cyclin D1 and c-MYC are well-known target genes of the β-catenin signaling pathway and are involved in oncogenic transformation [37]. Similarly, matrix metalloproteinase 2 (MMP-2) has been shown to play a vital role in tumor progression by regulating the tumor microenvironment, primarily by maintaining the connection between the tumor and stroma [38]. We observed profound increases in the expression of c-MYC, Cyclin D1, and MMP2 and the reversal of the LPA-induced effect after the addition of an LPAR2 receptor antagonist. Indeed, our most exciting observation was the significant overexpression of β-catenin and its downstream target genes in the clinical cohort studies of 415 human gastric cancer tissue samples, which is consistent with our data. These results confirm that LPA mediates its action in the initiation and progression of gastric cancer through the β-catenin signaling pathway and the LPAR2 receptor.The metabolic pathway used by cancer cells differs from that of normal cells. Normal cells use mitochondrial OXPHOS as a major source of energy, whereas the metabolic phenotype of each cancer is different due to its wide range of heterogeneity [39]. When cancer cells encounter unfavorable conditions, they change their metabolic phenotype to adapt themselves to the new microenvironment [40]. In the 1920s, the German scientist Otto Warburg established that cancer cells use aerobic glycolysis because mitochondrial OXPHOS is impaired [41,42,43]. Several factors are responsible for the induction of aerobic glycolysis, including the loss of tumor suppressor genes and the activation of oncogenes, the hypoxic microenvironment, the mutation of mitochondrial DNA, and the origin of the tissues [44]. Later, several other research groups demonstrated that mitochondrial OXPHOS is intact in many cancers [45,46]. It has also been reported that rather than the impairment of OXPHOS, the Warburg effect in cancer occurs due to increased glycolysis, which leads to the suppression of OXPHOS; when glycolysis is inhibited, OXPHOS is re-established. Although glycolysis is the major pathway of the energy metabolism in cancer, some cancers use OXPHOS or a mixture of OXPHOS and glycolysis [47,48]. Our findings suggest that gastric cancer cells exposed to LPA experience increased mitochondrial bioenergetics by increasing OXPHOS and glycolysis. It has been shown that during the early stage of carcinogenesis, cancer cells use glycolysis to fulfill the energy demands due to the oncogenic transformation and hypoxic conditions. However, when the cells undergo aglycemic conditions and a shortage of nutrients due to an increased proliferation and migration of signaling proteins into the nucleus, the cells use OXPHOS as a source of energy [48]. Previous reports have suggested that cancer cells use both OXPHOS and glycolysis because when cancer cells grow rapidly, they need more energy and metabolic intermediates for the biosynthesis of macromolecules. Many intermediates of glycolysis and a truncated TCA cycle can be used to synthesize the macromolecules required for the rapid growth and proliferation of cancer cells [49]. In 2009, Dang et al. showed that the ectopic expression of MYC in cancers can drive aerobic glycolysis and/or OXPHOS depending on the tumor cell microenvironment. MYC increases glycolysis by increasing the glycolytic genes; however, it can increase OXPHOS through the activation of O2-dependent glutaminolysis which results from increases in glutaminase due to the reduced expression of miR23a/b [50]. In addition, as a result of glycolysis, more lactate is produced; this generates an acidic tumor microenvironment that helps in the invasion and metastasis of cancer cells [51]. Again, lactic acidosis favors OXPHOS by inhibiting glycolysis [40]. Since the results from our functional and biochemical experiments showed that LPA increases proliferation and the subsequent progression of a gastric cancer cell line, and that MYC expression is increased after an LPA treatment, there is a possibility that the LPA increased glycolysis and OXPHOS due to the increased MYC expression. Shin et al. showed that Axin is present in the mitochondria of HeLa cells; when it is present in the mitochondria, it causes instability of the mitochondrial complex IV, resulting in the reduction in OXPHOS and ATP synthesis [52]. In addition, Park and colleagues also demonstrated a reduced Axin1 expression in an E-cadherin-overexpressed AGS cell line that underwent EMT and increased OXPHOS as a source of energy [53]. Our study also found that an LPA treatment reduced the mRNA level of Axin2 which could be another possibility for the increased mitochondrial energy metabolism. Our results also suggested that LPA-induced OXPHOS and the aerobic glycolysis of cells were significantly reduced following a treatment with the LPAR2 antagonist and XAV393 (Axin stabilizer). These results confirmed that LPA mediates its action through the LPAR2 receptor during the malignant progression of gastric cancer cells by increasing the energy metabolism due to the dysregulation of Axin2. However, how LPA-LPAR2 and dysregulated Axin2 induce glycolysis and OXPHOS in the gastric cancer cell line was not clearly shown in this study, suggesting that a further investigation is required to reveal a more detailed molecular mechanism.

Taken together, all of our results demonstrate a novel role for LPA via its LPAR2 receptor in the malignant progression of gastric cancer by activating the β-catenin signaling pathway and by the alteration of the energy metabolism. We demonstrated that LPA-LPAR2 increased ATP production by both OXPHOS and glycolysis via the downregulation of Axin2, thereby providing energy sources for the proliferation, migration, and invasion of gastric cancer cells. Thus, targeting the specific LPAR2 receptor and Axin2 may provide a novel therapeutic approach for the treatment of gastric cancer.