Peroxisome proliferator-activated receptor delta (Ppard) activation has been implicated in regulating a multitude of biological processes in placental development. In this study, we employed the UPLC-ESI-TOFMS approach to investigate the metabolic traits in placenta from GW501516-treated mice at Embryonic Day 10.5. By analyzing the mass spectrum data, ions with the most significant differences between control and GW501516-treated groups were identified. Among these metabolites, the fatty acids, phospholipids, and sterol lipids were dramatically increased. Ingenuity Pathway Analysis (IPA) showed that phosphatidylethanolamine biosynthesis and glycolysis were the top two altered metabolic pathways involved in carbohydrate metabolism, energy production, and lipid metabolism. Subsequent immunoblotting experiments provided evidence for positive correlation of PPARD level and AKT and ERK signaling pathways upon GW501516 treatment. Furthermore, the stimulation of GW501516 increased trophoblast cell fusion gene syncytin-A (Syna), but not syncytin-B (Synb), expression, suggesting a potential role of Ppard activation in promoting cytotrophoblast differentiation. Our results reveal that Ppard activation elicits dramatic changes of the metabolic activities in placenta, which is correlated to AKT and ERK signaling.
INTRODUCTION
The placenta is the first fetal organ to form during mammalian embryogenesis [12–3]. During gestation, the major function of placenta is to produce hormones, synthesize and release fatty acid, and facilitate efficient nutrient uptake [45–6]. A number of transcription factors have been implicated in placental development [78–9]; however, it remains unclear which factor is essential for this biological process.
Peroxisome proliferator-activated receptors (PPARs) are a family of the nuclear hormone receptors belonging to the steroid receptor superfamily. PPARs function as gene transcription factors in controlling lipid metabolism, energy homeostasis, and cell differentiation. The three subtypes have been described as Pparα, Ppard, and Pparg [10]. Each PPAR isotype has distinct tissue distributions, physiological functions, and ligands.
Among these three isotypes, Ppard is the most important regulator for executing key cellular functions in heart, liver, colon, and skeletal muscle. Physically, Ppard is activated by a large spectrum of endogenous and synthetic ligands, such as fatty acids, eicosanoids, prostacyclin, and GW501516 [1112–13]. Ppard activation elicits multiple metabolic responses found in vitro and in vivo, which include improving lipid oxidation and glucose uptake in skeletal muscle [14], stimulating lipid accumulation in keratinocytes [15], and increasing transintestinal cholesterol efflux [16]. Ppard activation has also been involved in cell differentiation [17], proliferation [18], and inflammation [19]. As a result, Ppard has been recognized as a major regulator in determining cell fate and a promising target in treatment of metabolic diseases [20].
In placenta, Ppard is the most ubiquitously expressed PPAR subtype in the early stage of pregnancy [21]. In particular, PPARD is more pronounced in the syncytiotrophoblast of human placenta [22]. A number of studies in Ppard-null mice have revealed that Ppard is involved in several aspects of placentation processes, including blastocyst implantation and decidualization, and is also critical for the integrity of the placenta-decidua interface [23, 24]. Loss of Ppard in mice confers placental defects, leading to more than 90% embryonic lethality between Embryonic Day (E) 9.5 and E10.5 [25], whereas activation of Ppard triggers massive trophoblastic giant cell differentiation with increased ADRP expression [26]. This implies that Ppard is a key regulator in placental metabolism and endocrine activities. PPARD agonist exposure has also been reported to negatively regulate phospholipids, cholesteryl esters, and cholesterol concentrations in normal human placenta [27, 28].
Considering the majority of placental Ppard-dependent metabolites have not been described as Ppard targets in placenta, the present study undertakes a metabolomics approach aimed to elucidate the specific metabolic and molecular pathways of Ppard activation in placental development upon Ppard agonist treatment.
MATERIALS AND METHODS
All protocols were approved by the Institutional Animal Care and Use Committee at Nanjing Medical University.
Animal
Female FVB mice were fed with the standard lab chow diet (3.1 kcal/g, containing 58% carbohydrate, 25% protein, and 12% fat; the diet was formulated in the laboratory) and water after weaning. Pregnancy was produced at 6 wk of age by overnight caging of a fertile female with a fertile male mouse. The next morning, the presence of a vaginal plug was defined as E0.5. Pregnant mice were randomized in two groups. Mice were orally administered vehicle (0.5% w/v aqueous methylcellulose solution) or a selective Ppard activator, GW501516 (kindly provided by Dr. Xirong Guo, Nanjing Medical University), at a dose of 2 mg/kg/day. Cesarean section was performed to collect both groups of placenta at E10.5 (n = 6 per group), E14.5 (n = 4 per group), and E18.5 (n = 4 per group). Samples were either frozen in liquid nitrogen or fixed in 10% formalin for further analysis.
Histopathology and Immunohistology
To investigate whether daily oral administration of GW501516 leads to morphological changes, placentas were collected at E10.5, E14.5 and E18.5, then sectioned and sequentially analyzed by histology. Immunohistochemistry was carried out on sections using the avidin-biotin-peroxidase method (Vector Laboratories) on 7-μm sagittal placental Paraffin-embedded sections. Endogenous peroxidases were quenched by exposing the sections to 3% H2O2 for 5 min, followed by a wash in PBS. Nonspecific binding sites were blocked by using 10% normal goat serum with 0.1% Triton-100 in PBS for 1 h. The sections were incubated with the primary antibody (see Supplemental Table S1; supplemental data are available online at www.biolreprod.org) overnight at 4°C. Slides were washed with PBS and incubated with appropriate biotinylated secondary antibodies (1:200; Vector Laboratories) for 45 min, followed by 30-min incubation in an avidin-biotin complex solution (Vector Laboratories). After washing three times in PBS, the sections were stained with peroxidase substrate solution until the desired stain intensity developed and then counterstained with hematoxylin solution for 5 min. Sections were mounted in Permount mounting media. The negative-control slides (immunoglobulin G only) were included in every individual analysis. Six mice in each group were used for quantification. Three high-power fields (400×) were randomly chosen for counting PPARD-immunostained cells in each sample. Approximately 1000 cells per group were analyzed. Student t-test was used to determine the level of significance (P < 0.05) between two groups. Data are presented as the mean ± SD.
Western Blot Analysis
The placentas were frozen in liquid nitrogen and pulverized in a mortar and pestle, then mixed with lysis buffer containing 50 mM Tris-HCl (pH 7.5), 0.5% NP-40, 0.1% SDS, 0.25% sodium deoxycholate, 125 mM NaCl, 1 mM ethylenediaminetetra-acetic acid, 50 mM NaF, 1 mM sodium orthovanadate, 2.5 mM sodium pyrophosphate, 1 mM sodium β-glycerophosphate, 1 mM PMSF, and protease inhibitor cocktail (Roche Diagnostics). Following incubation on ice for 30 min, the lysates were cleared by centrifugation at 13 000 × g for 15 min at 4°C. Protein concentrations were determined by Coomassie Plus Protein Assay (Pierce), and then 50 μg of lysate were separated in a 4%–12% NuPAGE Bis-Tris gel (Invitrogen). After wet transfer, membranes were blocked for 1 h at room temperature in Tris-buffered saline (pH 7.4) containing 5% nonfat dry milk and 0.1% Tween 20, then incubated with appropriate primary antibody (see Supplemental Table S1) overnight at 4°C and horseradish peroxidase-linked secondary antibodies for 1 h at room temperature. Proteins were visualized with either SuperSignal West Pico or SuperSignal West Dura (Pierce). The band intensity was quantified by Scion Image software. Band intensity was normalized to total protein or actin in respective columns. Wilcoxon rank-sum test was used to determine the level of significance (P < 0.05) between two groups. Results are presented as the mean ± SD from three separate experiments.
Sample Preparation and Mass Spectral Analysis
To gain further understanding of the effect of GW501516, we measured the mouse placental metabolism at E10.5 by using the ultraperformance liquid chromatography (UPLC) and the accurate mass measurement of electrospray ionization time-of-flight mass spectrometry (ESI-TOFMS) approach. Cold methanol extracts of placentas from six controls and an equal number of mice treated with GW501516 for 10 days were applied for liquid chromatography (LC)-MS analysis. Samples were clarified and processed as described below: Each sample (5 μl) was injected onto a reverse-phase 50- × 2.1-mm ACQUITY 1.7-μm C18 column (Waters Corp.) using the ACQUITY UPLC System (Waters) with a gradient mobile phase consisting of 2% acetonitrile in water containing 0.1% formic acid (A) or 2% water in acetonitrile containing 0.1% formic acid (B). Each sample was resolved for 10 min at a flow rate of 0.5 ml/min. The gradient consisted of 100% A for 0.5 min, then a ramp of curve 6 to 100% B from 0.5 to 10 min. The column eluent was introduced directly into the mass spectrometer by electrospray. The desolation gas flow was set to 800 L/h, and the temperature was set to 350°C. The cone gas flow was 25 L/h, with a source temperature of 120°C. Accurate mass was maintained by introduction of LockSpray interface of leucine-enkephalin (556.2771 [M+H]+ or 554.2615 [M-H]−) at a concentration of 2 ng/μl in 50% aqueous ACN with an infusion rate of 2 μl/min for the positive mode and 3 μl/min for the negative mode.
Metabolites Identification and Data Preprocessing
The UPLC-ESI-TOFMS data in positive and negative modes were analyzed by the MassLynx software (Waters) with the format of Network Common Data Form. Then, XCMS Online [29] was used for candidate identification. The parameter was optimized with UPLC/Q-TOF high-resolution LC-MS data according to Patti et al. [30]. The ions with a fold-change larger than 1.5 and a P-value less than 0.05 were selected for further analysis. Total ion chromatograms plot, mirror plot, and principal components analysis results were provided by XCMS Online. Boxplot and extracted-ion chromatogram peaks were also generated by XCMS Online and sorted in the order of most to least significant. Putative identifications for the resulting ion list were obtained through a database search tool [31] that combines the following four metabolite identification databases (Human Metabolite DataBase [32], Metlin [33], Madison Metabolomics Consortium Database [34], and LIPID MAPS [35]) with mass tolerance set to 10 ppm. For metabolite statistical analysis, R scripts developed in-house were used to select the ions with significant and consistent changes between the treated and control groups. To delineate the early metabolic consequences of Ppard activation, metabolomics analysis was carried out with extracts (n = 6 per group) from control and GW501516-treated placenta based on one-way ANOVA and the variability of the corresponding peaks in the quality control runs of the placental samples.
Quantitative Real-Time PCR
Total RNA was extracted using TRIzol (Sigma) from E10.5 placentas following the manufacturer's protocol. One microgram of RNA was reverse transcribed in a total volume of 20 μl using the Omniscript Reverse Transcription Kit (Qiagen). Quantitative real-time PCR (qPCR) was performed with the primers CTGGGAATATGAACCCACTGTTA and GAGTTGAGGCAGAAGGAGTATG for syncytin-A (Syna) and the primers CCACCACCCATACGTTCAAA and GGTTATAGCAGGTGCCGAAG for syncytin-B (Synb). The qPCRs were performed in triplicate in an ABI Prism 7700 Instrument (Applied Biosystems) with QuantiTect SYBR Green (Qiagen) following the manufacturer's protocol. The PCR products were confirmed by ethidium bromide staining on a 2% agarose gel. The expression of each target gene was presented as the ratio of the target gene to 18S RNA, which was expressed as 2−ΔCt, where Ct is the threshold cycle and ΔCt = CtTarget − Ct18S. Students t-test was applied to determine the level of significance (P < 0.05) between two groups. Values represent the mean ± SD from three separate experiments.
Molecular Pathway and Network Analysis by Ingenuity Pathway Analysis
Ingenuity Pathway Analysis (IPA) was applied to identify biological pathways and functions relevant to biomolecules of interest. A ratio of metabolites that map to the pathways divided by the total number of molecules that map to canonical pathways was displayed. Fisher exact test was adapted to calculate the P-value determining the probability that the association between the metabolites and the canonical pathway was explained by chance alone. The network score was based on the hypergeometric distribution and was calculated by two-tailed Fisher exact test. The higher the score, the more relevant the eligible submitted molecules were to the network.
RESULTS
Histologic Analysis of Ppard Agonist Effects on Mouse Placenta
Histologic analysis indicated no significant placental morphologic change between the GW501516-treated group and the control group at E10.5 (Supplemental Fig. S1) or at E14.5 and E18.5 (data not shown). We investigated the temporal/spatial pattern of PPARD expression in mouse placenta of E10.5. PPARD was predominantly expressed in syncytiotrophoblast in the labyrinth zone. GW501516-treated placenta showed a greater number of PPARD-positive cells than control placenta (P < 0.01) (Fig. 1A). Western blots were performed for PPAR protein expression analysis on placentas at E10.5, E14.5, and E18.5. GW501516 treatment induced detectable increase of endogenous PPARD protein expression, whereas PPARA and PPARG levels were not affected by GW501516 (Fig. 1B). PPARD protein levels were significantly increased in GW501516-treated mice at E10.5 (P < 0.01) (Fig. 1C).
FIG. 1
The effects of Ppard agonist on mouse placenta. A) Immunohistochemical analysis indicated that PPARD is expressed in syncytiotrophoblast cells and increased in response to GW501516. Representative images from control and GW501516-treated mice are shown. Bar graph indicates the number of PPARD-positive cells per 1000 cells based on a total of six samples. **P < 0.01, Student t-test. Magnification ×400. B and C) Representative Western blots (B) and quantification (C) of PPARA, PPARG, and PPARD protein expression in mice from E10.5 (n = 6), E14.5 (n = 4), and E18.5 (n = 5) placenta. All values are normalized to actin. Each value represents the mean ± SD, expressed relative to control values. Student t-test was performed on the data. **P < 0.01, *P < 0.05 denotes significant differences between GW501516-treated and control groups.
Analysis of Global Metabolic Response of Placenta to Ppard Agonist
To evaluate if the UPLC-ESI-TOFMS-based metabolomics approach is useful to discriminate GW501516-treated placentas from controls (n = 6 per group), placental extracts were profiled using UPLC-ESI-TOFMS in positive mode and native mode, with the data preprocessed by XCMS. We detected 531 m/z chemically identified monoisotopic ion masses with significant difference in the treated group versus the control group (fold-change ≥ 1.5, P < 0.05). The base peak plot (Fig. 2A), mirror plot (Fig. 2B), and heatmap visualization (Fig. 2C) showed distinct segregation between GW501516 treatment and controls.
FIG. 2
UPLC-ESI-TOF-MS profiling of metabolites altered in placenta. In all, 561 metabolites for which expression was altered more than 1.5-fold were applied to IPA. A) Base peak plot shows the overlay of total ion chromatograms of GW501516-treated group (red) and control group (black). B) Mirror plot of placenta differential metabolites. Up-regulated features are shown in green and down-regulated features in red. C) Heat map depicts the changes of metabolites for control and GW501516-treated placenta (n = 6 individual measurements per group).
Identification of Metabolites Affected by Ppard Agonist
Table 1 presents the most relevant metabolites identified from these monoisotopic ion masses in distinguishing the treated group from the control group. The change of a metabolite is indicated with an average fold-change value. In all, 59 of 67 metabolites listed in Table 1 were up-regulated, and 8 of 67 metabolites were down-regulated, in response to GW501516. The metabolites of GW501516-treated mice, in comparison to those of control mice, exhibited an increase in fatty acid and phospholipid levels of eicosatrienoic acid (11.1-fold), (all Z)-7,10,13,16,19-docosapentaenoic acid (DPA; 16.6-fold), eicosapentaenoic acid (EPA; 16.6-fold), phosphatidylcholine (7.1-fold), phosphatidylethanolamine (PE; 7.1-fold), elaidic acid (8.0-fold), and arachidonic acid (AA; 5.674-fold), whereas the levels of l-threonine (−1.8 fold), sebacic acid (−1.6 fold), magnolol (−2.79 fold), and honokiol (−2.79 fold) were decreased in response to GW501516. As shown in Figure 3, an overall change in lipid composition in response to GW501516 was the significant increase of fatty acids, phospholipids, and sterol lipids.
TABLE 1
The most relevant metabolites identified in the GW501516-treated group compared to control group.
a
KEGG, Kyoto Encyclopedia of Genes and Genomes.
Metabolic Pathway and Function Analysis
The 67 identified and quantified metabolites were applied to IPA. Figure 4 presents the top canonical pathways enriched in the metabolites identified with GW501516 in placenta, including PE biosynthesis and glycolysis. As shown in Table 2, network analysis of 67 differentially expressed metabolites revealed three significantly altered metabolism networks, including the key metabolites DPA, AA, PE, and 15(S)-HETE (Table 2). These metabolic networks were mainly involved in the following metabolite subset functions: 1) carbohydrate metabolism, energy production, and endocrine system development and function; 2) cell signaling and vitamin and mineral metabolism; and 3) lipid metabolism, small molecule biochemistry, and cellular assembly and organization.
FIG. 4
Metabolites from the GW501516-treated group at E10.5 were associated with 33 distinctly different canonical pathways identified by IPA (P < 0.05). The y-axis represents the –log of P-values calculated by Fischer exact test. A threshold of P = 0.05 is represented by the light yellow line. This gives a measure for the possible impact of GW501516 on each listed pathway. The orange line represents the proportion of the amount of metabolites involved in some pathways.
TABLE 2
IPA for metabolites in top related network and functions are listed.
Role of Ppard Activation on AKT and ERK Cell Signaling Pathways
The mechanistic network function analysis of IPA showed that AKT and ERK signaling pathways were predicted to be activated in response to GW501516 (Fig. 5). For further verification, Western blot analysis was conducted with mouse placenta tissues. As shown in Figure 6A, GW501516 treatment dramatically increased pTh308AKT (*P < 0.05), pSer473AKT (*P < 0.05), and pERK1/2 (*P < 0.05) levels at E10.5 compared to the untreated group (Fig. 6B).
FIG. 5
Metabolite expression regulatory networks predicted by IPA based on the metabolites listed in Table 1. AKT and ERK were predicted to be activated. Each node represents a metabolite. Red nodes denote up-regulated metabolites, and green nodes denote down-regulated metabolites. Dotted lines indicated indirect regulatory interactions, whereas solid lines show direct regulatory interactions.
FIG. 6
The effects of Ppard agonist on AKT and ERK signaling pathways. A) Representative Western blots of the signaling pathways underlying Ppard activation in placentas. GW501516 led to significant increase in Th308AKT, Ser473AKT, and ERK phosphorylation. B) Bar graph of the ratio of pTh308AKT/AKT, pSer473AKT/AKT, and pERK1/2/ERK1/2 shown in A. All values are normalized to actin. Each value represents the mean ± SD, expressed relative to control values. Student t-test was performed on the data. **P < 0.01, *P < 0.05 denotes significant differences between GW501516-treated versus control groups.
Role of Ppard Activation on Placental Syn Expression
Syna and Synb were used to evaluate the effect of Ppard activation on placental function. Syna, but not Synb, was significantly increased (P < 0.01) in the GW501516-treated group compared to the control group (Fig. 7). This indicates that Ppard activation may enhance cytotrophoblast differentiation by inducing Syna expression.
FIG. 7
The effects of Ppard agonist on syncytin expression. The qPCR analysis of the placental cytotrophoblast differentiation markers syna and synb gene expression is shown. Syna significantly increased in GW501516-treated placenta in comparison to control mice, whereas no significant change was found in Synb expression (n = 6 mice/group) between the two groups. The data represent the mean ± SD. **P < 0.01 denotes significant differences between GW501516-treated versus control groups.
DISCUSSION
The placenta is a prominent organ for glucose transport, fatty acid synthesis, and uptake for fetal growth and development. The metabolic activities in placenta are important for nutrient transport [36, 37]. In the present study, we characterized the metabolic changes pertaining to Ppard activation using the UPLC-ESI-TOFMS approach and function enrichment analysis. Our results indicate that placenta treated with GW501516 does not exhibit phenotype changes throughout pregnancy, which is different from the results of Nishimura et al. [38]. The use of different dosages of GW501516 (2 vs. 100 mg/kg/day) and different models (mice vs. rats) may have contributed to this difference.
The temporal/spatial expression patterns of PPARD suggest that oral GW501516 administration may have a direct influence on placenta. Our PPAR expression results indicate that PPARA and PPARG expression was not altered by GW501516, because GW501516 is the most potent PPARD agonist known, with a median effective concentration of 1 nM. Moreover, it has higher selectivity (>1000-fold) over the other two PPAR isoforms [39]. We speculate that the actions of PPARD agonist on placenta likely are PPARD-dependent in the present study.
We demonstrated that Ppard activation alters the content and composition of placental lipids. Of the 67 most relevant metabolites, 34 consist of fatty acids. In particular, a significant increase of the main components of polyunsaturated fatty acids was observed, including DPA, EPA, and AA in GW501516-treated groups versus controls. DPA is an intermediate product between EPA and docosahexaenoic acid (DHA). AA and DHA have been implicated in proper fetal brain and eye development [40]. EPA is positively correlated with fatty acid binding protein mRNA expression [41] that, in turn, enables free fatty acid, such as DHA and AA, to cross the placenta to meet the demands of the normal growing fetus, whereas insufficient intake of EPA during pregnancy is associated with preterm birth [42]. Thus, the changes in placental lipid contents caused by GW501516 appear to be involved in the modulation of free fatty acid transport processes.
Previous studies have reported that activation of Ppard not only induced gene expression involved in energy production in adipose tissue and skeletal muscle [43, 44] but also significantly improved glucose tolerance and insulin resistance in the animal model of high fat diet-induced obesity [45]. On the other hand, loss of Ppard in mouse muscle exhibited reduced oxidative and metabolic efficiency [46]. These studies emphasize the regulatory role of Ppard in fatty acid oxidation, energy expenditure, and glucose homeostasis. However, the role of Ppard pertaining to placental metabolic activates is not well clarified. Our study revealed that the network most altered by GW501516 involved a highly interconnected carbohydrate and energy metabolism, characterized by up-regulated glycolysis and lipid biosynthesis activities. These results, in agreement with previous reports on the beneficial effects of Ppard on lipid metabolism and whole-body glucose tolerance [434445–46], implicates that activation of Ppard may offer an effective strategy in preventing gestational diabetes by increasing placental capacity for fatty acid oxidation, energy expenditure, and glucose homeostasis. Given the loss of Ppard in mice causes induction of embryo lethality as well as reduction of energy uncoupling and adipose mass [25], such effects on placental metabolic activities may also explain the placental malformation outcomes of Ppard deficiency.
In addition to its role in transporting molecules between mother and fetus [47], the placenta is a major endocrine organ that provides abundant hormones for mammalian reproductive success [48]. Our finding that Ppard activation leads to the augmented sterol lipids, such as desoxycorticosterone acetate (4-fold change) and cholesterol sulfate (3.4-fold change), indicates the involvement of Ppard activation in prompting the placental endocrine secretions, which are crucial for placental development and fetal growth. Interestingly, the metabolism analysis showed the positive effects of GW501516 on AA (5.7-fold change), because AA per se is one of the endogenous Ppard ligands that not only regulates human chorionic gonadotropin-induced steroidogenesis but also plays a role in testosterone formation in rat Leydig cells [49, 50]. Thus, the increased AA in the present study may explain why Ppard activation confers placental endocrine system development and placentation.
Furthermore, our analysis of biology function predicted that Ppard might positively regulate AKT and ERK signaling pathways mediated by metabolites during pregnancy. In agreement with the network analysis from IPA, the Western blot analysis showed that Ppard activation resulted in an increase of pTh308AKT, pSer473AKT, and pERK1/2 expression. Although the direct mechanical link between Ppard activation and AKT/ERK signaling is missing, our metabolomics profiling suggests that Ppard activation leads to elevated levels of fatty acids, which presumably drive the activation of the AKT indirectly through the PI3K/PDK1/AKT axis [51] and the phosphorylation of ERK through the EGFR/ERK/p90RSK pathway [52]. Notably, 15(S)-HETE is a downstream product of AA and enhances the expression of placental growth factor through PI3/AKT signaling pathway [53]. Moreover, the interaction between Ppard and PI3K/AKT signaling was found in other cell types [54, 55]. Given that Ppard is a fatty acid sensor in regulating lipid metabolism in several metabolic tissues [56], the present study provides a molecular mechanical linking between Ppard and AKT/ERK activities in the placenta metabolic processes.
Syncytin, a fusogenic glycoprotein of endogenous retroviral origin, plays a critical role in the regulation of cytotrophoblast differentiation [57]. Among the PPAR family, the role of PPARG in regulating syncytin-1 mRNA expression as well as trophoblast differentiation and invasion is well established [58, 59]. However, Ppard effects on cytotrophoblast differentiation in placentas are unknown. Our data show that GW501516 increased Syna, but not Synb, mRNA expression, suggesting that Ppard activation positively regulates cytotrophoblast differentiation. Because the formation of syncytium coincided with glycolysis and energy production during cytotrophoblast differentiation [60], we speculate that the role of Ppard activation is regulating Syna expression via the modulation of metabolic pathway, which was reflected by induced glycolysis upon GW501516 treatment. In addition, as the 5′-long terminal repeat of human endogenous retrovirus-W (HERV-W) promoter contains three putative PPAR response elements (PPREs 1–3), which suggests that Ppard may directly regulate Syn transcription [59]. Thus, the effect of Ppard activation in promoting cytotrophoblasts differentiation merits further attention.
In summary, our data suggest that Ppard activation is associated with changes of lipid content in the placenta that correspond with unregulated AKT/ERK signaling pathways and Syna expression. Further studies in human placenta are needed to better understand the impact of altered metabolic functions of Ppard activation on placental development, fetal growth, and the cellular mechanisms involved.
ACKNOWLEDGMENT
We are grateful to Dr. Xirong Guo for help with the present study. We thank Dr. Jinlian Wang for data analysis.
