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Differentiation of Trophoblast Giant Cells and Their Metabolic Functions Are Dependent on Peroxisome Proliferator-Activated Receptor ß/

Karim Nadra,¹ Silvia I. Anghel,^1,2 Elisabeth Joye,¹ Nguan Soon Tan,^1, Sharmila Basu-Modak,^1, Didier Trono,^2,3 Walter Wahli,^1,2 and Béatrice Desvergne^1,2*

Center for Integrative Genomics, University of Lausanne, CH-1015 Lausanne, Switzerland,¹ National Research Center "Frontiers in Genetics"^, , School of Life Sciences, Swiss Federal Institute of Technology, Lausanne, Switzerland³

Received 24 June 2005/ Returned for modification 21 July 2005/ Accepted 24 January 2006

	ABSTRACT

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Mutation of the nuclear receptor peroxisome proliferator-activated receptor ß/ (PPARß/) severely affects placenta development, leading to embryonic death at embryonic day 9.5 (E9.5) to E10.5 of most, but not all, PPARß/-null mutant embryos. While very little is known at present about the pathway governed by PPARß/ in the developing placenta, this paper demonstrates that the main alteration of the placenta of PPARß/-null embryos is found in the giant cell layer. PPARß/ activity is in fact essential for the differentiation of the Rcho-1 cells in giant cells, as shown by the severe inhibition of differentiation once PPARß/ is silenced. Conversely, exposure of Rcho-1 cells to a PPARß/ agonist triggers a massive differentiation via increased expression of 3-phosphoinositide-dependent kinase 1 and integrin-linked kinase and subsequent phosphorylation of Akt. The links between PPARß/ activity in giant cells and its role on Akt activity are further strengthened by the remarkable pattern of phospho-Akt expression in vivo at E9.5, specifically in the nucleus of the giant cells. In addition to this phosphatidylinositol 3-kinase/Akt main pathway, PPARß/ also induced giant cell differentiation via increased expression of I-mfa, an inhibitor of Mash-2 activity. Finally, giant cell differentiation at E9.5 is accompanied by a PPARß/-dependent accumulation of lipid droplets and an increased expression of the adipose differentiation-related protein (also called adipophilin), which may participate to lipid metabolism and/or steroidogenesis. Altogether, this important role of PPARß/ in placenta development and giant cell differentiation should be considered when contemplating the potency of PPARß/ agonist as therapeutic agents of broad application.

	INTRODUCTION

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The earliest cell differentiation event in the mammalian embryo is the formation of the trophectoderm, which consists of two regions: the polar trophectoderm, directly adjacent to the inner cell mass, and the mural trophectoderm surrounding the blastocoelic cavity. In rodent placentas, the cells of the mural trophectoderm differentiate into a limited number of nondividing primary giant cells during implantation, whereas those of the polar trophectoderm proliferate to form the ectoplacental cone and develop a multitude of secondary giant cells (20).

Trophoblast giant cells participate in a number of processes essential to a successful pregnancy, including blastocyst implantation, remodeling of the maternal deciduas, and secretion of hormones that regulate the development of both the fetal and maternal compartments of the placenta. In the early stages, the inherent invasive properties of these cells appear to be crucial for remodeling in the maternal uterine stroma. At later stages, after embryonic day 9.5 (E9.5), the secondary giant cells produce a number of placental hormones, particularly members of the prolactin/growth hormone family of proteins, which in mice includes placental lactogen I (PL-I), PL-II, and proliferin (49). The expression of these trophoblast giant cell functions coincides with a very critical period of placental development, exemplified by the high incidence of embryonic lethality at that stage. Paradoxically, very little is yet known about the giant cell differentiation process.

Peroxisome proliferator-activated receptors (PPARs) are lipid-activated transcription factors that belong to the nuclear hormone receptor family. Three isotypes of PPARs have been cloned: PPAR (NR1C1; Nuclear Receptor Nomenclature Committee 1999), PPARß/ (NR1C2; also called NUC-1 or FAAR and which will be hereafter referred to as PPARß), and PPAR (NR1C3), all of which bind to DNA as heterodimers with retinoid X receptor (RXR; NR2B) (10). While PPAR and PPAR have clearly defined roles in controlling lipid and glucose homeostasis (64), various physiological roles of PPARß are still being investigated. PPARß has been linked to embryo implantation (41, 42), myelination in the brain (51), osteoclastic bone resorption (44), and skin wound healing (46, 58). Its role in lipid metabolism comprises diverse facets associated with the preadipocyte clonal expansion (26, 29), fatty acid oxidation in muscle (63), or lipoprotein homeostasis (50). The nature of the endogenous PPARß ligands remains to be ascertained. Similar to the other two PPARs, some polyunsaturated fatty acids have affinities for PPARß in the low micromolar range. More specifically, a number of eicosanoids, particularly prostacyclin, which is a cyclooxygenase 2 arachidonate metabolite, was shown to activate PPARß (16, 35, 37).

A crucial tool for understanding PPAR functions in an in vivo and physiological context is the generation of strains of mice carrying mutations in the PPAR genes. However, most of the PPARß and all PPAR null embryos die at an early developmental stage because of placental defects (1, 2, 38) that have so far been poorly characterized. In the present work, we delineate a pivotal role of PPARß in the early events of placental development. Whereas deficiency of PPAR leads to impaired vascularization (1, 38), deletion of the PPARß gene causes a severe failure of the placenta to undergo proper morphogenesis, leading to embryonic lethality at E9.5 to E10.5. The trophoblast giant cell layer is the most affected, and we demonstrate a direct involvement of PPARß in promoting trophoblast cell differentiation toward giant cells. This effect is dependent on phosphatidylinositol 3-kinase (PI3K) and Akt1 (also called protein kinase B) and is at least partly due to the high expression levels of two kinases involved in Akt activation, namely 3-phosphoinositide-dependent kinase 1 (PDK1) and integrin-linked kinase (ILK). In addition, PPARß caused increased expression of a non-helix-loop-helix (non-HLH) inhibitor of the myogenic basic HLH (bHLH) subfamily, I-mfa (7), which contributes to the differentiation process. Finally, we reveal that giant cells are the primary sites of lipid accumulation in the placenta at an early stage, together with a PPARß-dependent up-regulation of adipose differentiation-related protein (ADRP) expression. Taken together, these cellular and in vivo approaches unveil important new aspects of the development and functions of the giant cell layer, which has a major impact on placenta development.

	MATERIALS AND METHODS

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Targeted disruption of the mouse PPARß. (i) Construction of a replacement vector for PPARß. Nine overlapping mouse PPARß genomic clones were isolated from an Sv129 mouse embryonic stem (ES) cell genomic library (gift from F. Conquet), and their restriction maps were established (Fig. 1). A targeting vector was designed to delete the two exons encoding the DNA binding domain (i.e., exons 4 and 5), according to the organization of the Xenopus PPARß gene. The targeting vector (derived from TK-NEO-UMS, a vector comprised of the thymidine kinase gene, neomycin resistance sequence, and upstream mouse sequence; a gift from Charles Weissman [54]) contained 1.7 kb of homologous sequence at the 5' end, 7 kb of homologous sequence at the 3' end, and a phosphoglycerate kinase-neomycin (PGK-neo) cassette (Fig. 1A).

View larger version (28K): [in this window] [in a new window]   FIG. 1. Targeted disruption of the PPARß gene in mouse. The PPARß gene was disrupted in ES cells by homologous recombination with a replacement-type vector, using an approach based on positive-negative selection (A). In this vector, PPARß genomic sequences containing the exons encoding the DNA-binding domain of the receptor (exon 4 and part of exon 5) were replaced with a PGK-neo cassette. Homologous recombination at the PPARß locus in ES cells led to the deletion of both exon 4 and part of exon 5 encoding the two zinc fingers of the DNA-binding domain. ES cells carrying the mutant allele were confirmed by Southern blot analysis (B). Two independent positive ES cell clones were injected into blastocysts to generate chimeras, and heterozygous mice were obtained from a germ line transmitter chimera. Panel A shows the structure of the wt PPARß allele, targeting vector, and recombinant PPARß allele. The exons as well as locations of restriction sites and probes for PCR and southern blot are indicated. B, BamHI; E, EcoRI; K, KpnI; N, NotI; X, XhoI. Panel B shows a Southern blot analysis of genomic DNA digested with BamHI and KpnI from E9.5 embryos produced by a PPARß heterozygous intercross. (C) PCR analysis of yolk sac DNA derived from E9.5 embryos. (D) Western blot analysis performed on nuclear extracts with a specific PPARß antibody. The nuclear protein c-Jun was used as an internal control. In order to obtain a sufficient amount of material, the control at the protein level was performed on pups obtained from homozygous matings.

(iii) Genotyping. Genomic DNA was prepared from ES cells, yolk sacs of embryos, or tail samples following the classical procedures. A first round of genotyping was performed by two independent PCRs. Primers b19 (5'-ATCCAGAGTGTTCGTATGAC-3') and UMS1 (5'-TCTTATGCTCCTGAAGTCCAC-3') amplified a 2.2-kb fragment from the recombinant allele, whereas the primers b3 (5'-AGCCTCAACATGGAATGTCG-3') and b4 (5'-GATCGCACTTCTCATACTCG-3') amplified a 1.6-kb product from the wild-type (wt) allele. Five percent of the neomycin-resistant ES cell clones were positive for homologous recombination. All mutant clones and/or embryos or mice were subsequently confirmed by Southern analyses using a digoxigenin-labeled probe (CDP-Star protocol; Boehringer Mannheim), located 160 bp upstream of the 5' homology region. Digested genomic DNA samples were blotted on a Zetaprobe GT membrane and processed following the manufacturer's protocol (Bio-Rad). Probes, restriction digestion, and hybridized fragments from wt and recombinant alleles are indicated in Fig. 1.

(iv) Generation of chimeric mice and germ line transmission. Positive D3 clones were microinjected into the blastocoel of 3.5-day-old embryos at the blastocyst stage and isolated from C57BL/6 females (10 to 15 ES cells per blastocyst). Between five and seven injected blastocysts were reimplanted into each uterine horn of pseudopregnant foster mothers. Male chimeric animals were mated for germ line transmission with Sv129 mice. One chimeric male transmitted the mutation from which the colony of mice has been obtained.

Reagents and probes. The antibodies anti-Akt1, anti-PTEN, anti-ILK, phospho-Akt (Thr308), and (Ser473) were obtained from Cell Signaling; anti-c-Jun and anti-PPARß were from Affinity Bioreagents; anti-PDK1 was from Santa Cruz Biotechnology; antitubulin was from Pharmingen; anti-ADRP was from Progen; and anti-PL-I was from Chemicon.

The gene-specific probes for PL-I and Tpbpa were a kind gift of J. Rossant, and Mash-2 was obtained from F. Guillemot. For preparing the probes, total RNAs were obtained from E9.5 placenta. cDNAs were prepared by reverse transcription, followed by PCR, using specific primers (primer sequences are available upon request). The cDNA corresponding to the mouse PPARß and L27 were subcloned into pGEM3Zf(+) (Promega). The cDNA corresponding to the mouse PPAR, as well as gene-specific probes corresponding to PL-I, Id-2, Hand1, I-mfa, ADRP, and aP2, was subcloned into pGEM-T Easy (Promega). Gene-specific antisense and sense riboprobes were synthesized by in vitro transcription with either T7 or Sp6 RNA polymerase (Ambion).

RPA. For all riboprobes, except L27, a ratio of 1:1 of [-³²P]UTP to cold UTP was used, whereas a ratio 1:20 was used for L27 probe. An RNase protection assay (RPA) was carried out as described by the manufacturer (Ambion) with the following modifications. Briefly, total RNA was isolated from cells or tissues with Trizol reagent (Sigma, St. Louis, MO). Ten micrograms of total RNA was incubated at 37°C with 1 ng of gene-specific riboprobes (10⁹ cpm/µg) and 10 ng of the L27 probe (10⁷ cpm/µg). RNase digestion (10 U/ml RNase A; 400 U/ml RNase T₁) was carried out for all probes at 37°C for 20 min. After inactivation of RNase with 20 µl of 10% sodium dodecyl sulfate (SDS) and 10 µl of 20 mg/ml proteinase K for 20 min at 37°C, protected RNA was precipitated with 500 µl isopropanol, and RPA products were resolved in a 6% electrolyte-gradient denaturing polyacrylamide gel. Gels were then dried and exposed to X-ray film or to the phosphor screen of a StormImager 840 (Molecular Dynamics). Control RPAs were performed using an L27 antisense probe, which has been described previously (39). Quantitative analysis was performed by using IQuant 2.5 software.

In situ hybridization. Embryos and placentas were obtained by dissecting pregnant females, with the appearance of a vaginal plug being counted as day 0.5 of pregnancy. Because of their close contact, particularly at E9.5 stage, both the maternal decidua and the fetal part of the placenta were dissected as a whole. Embryos and placentas were separately fixed overnight at 4°C in 4% paraformaldehyde in phosphate-buffered saline (PBS) and embedded in paraffin, and 7-µm sections were cut. In situ hybridization was processed as described previously (5). Briefly, antisense and sense riboprobes for PPARß, Mash-2, Tpbpa (or 4311), and PL-I were generated by in vitro transcription of the corresponding cDNA clones with SP6 or T7 RNA polymerase. Rehydrated sections were washed in 2x SSC (1x SSC is 0.15 M NaCl plus 0.015 M sodium citrate) for 5 min, and hybridization was performed overnight at 70°C with 1 µg/ml of a digoxigenin-labeled probe in the following hybridization buffer: 50% formamide, 10% dextran sulfate, 1x Denhardt's solution, 1 mg/ml yeast RNA, 200 mM NaCl, 1.1 mM Tris-base, 8.9 mM Tris-HCl, 5 mM Na₂HPO₄, and 1.25 mM EDTA. Three washes of 30 min were done in 50% formamide, 1x SSC, and 0.1% Tween 20 at 65°C, followed by two washes in MABT (100 mM maleic acid, 140 mM NaCl, 1% Tween 20, pH 7.5). Sections were then incubated for 90 min in a blocking buffer consisting of 20% goat serum-2% blocking reagent (Boehringer Mannheim) in MABT and then left overnight at room temperature in an alkaline phosphatase-conjugated antidigoxigenin antibody (Boehringer Mannheim) at a dilution of 1:2,000 in a blocking buffer. Washes were done in MABT, five times for 30 min each time. Sections were then washed for 5 min in NTMT (100 mM NaCl, 100 mM Tris, 50 mM MgCl₂, 1% Tween 20, pH 9.5), and a color reaction assay was performed in the dark in nitroblue tetrazolium and 5-bromo-4-chloro-3-indolyl phosphate (Roche) and 1 mM levamisole in NTMT.

Immunohistochemistry. Immunohistochemistry was carried out on sections using the avidin-biotin-peroxidase method (Vector Laboratories) on 7-µm sagittal placental paraffin 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 using 10% normal serum in 0.1% Triton-PBS for 1 h. Immediately after, the sections were incubated with the primary antibody for 1 h. After being washed in PBS, the slides were incubated with appropriate biotinylated secondary antibodies (1:200; Vector Laboratories) for 45 min, followed by 30 min in an avidin-biotin complex solution (Vector Laboratories) in PBS. The sections were then stained with 3,3' diaminobenzidine (Sigma), rinsed in water, and counterstained with a 0.1% methyl-green aqueous solution for 5 min.

Western blots. For the Western blot of Fig. 1, nuclear extracts were obtained from the skin of mouse pups. Briefly, tissues were homogenized in 10 mM KCl, 1.5 mM MgCl₂ 10 mM HEPES, pH 7.9, 1 mM dithiothreitol, 1 mM Na₃VO₄, 10 µg/ml leupeptin, 0.3 trypsin inhibitor units (TIU)/ml aprotinin, 1 µM pepstatin A, and 1 mM phenylmethylsulfonyl fluoride. The homogenate was centrifuged at 2,000 x g for 10 min, and the pellet was washed with the same buffer. The washed pellet was resuspended in a buffer containing 420 mM NaCl, 10 mM KCl, 20 mM HEPES, pH 7.9, 20% glycerol, 1 mM dithiothreitol, 1 mM Na₃VO₄, and the protease inhibitors specified above.

For other experiments, Western blotting was performed on cells or tissues, lysed in an ice-cold lysis buffer (20 mM Na₂H₂PO₄, 250 mM NaCl, 1% Triton 100, 0.1% SDS) supplemented with complete protease inhibitors (Roche). After Bradford quantification (Bio-Rad), 10 µg of nuclear extracts or 30 µg of total protein was separated by SDS-polyacrylamide gel electrophoresis and transferred onto polyvinylidene difluoride membranes for Western blotting. The membranes were processed in the following steps: (i) 1 h of saturation in Tris-buffered saline (TBS)-0.1% Tween-5% nonfat milk at room temperature (RT), (ii) overnight incubation with a primary antibody in TBS-0.1% Tween-5% nonfat milk at 4°C, (iii) three washings in TBS-0.1% Tween at RT, (iv) exposure for 1 h to a secondary antibody in TBS-0.1% Tween-5% nonfat milk at RT, and (v) three washings in TBS-0.1% Tween at RT as a final step. The signal was detected using an ECL detection kit (Amersham Pharmacia Biotech), according to the manufacturer's instructions. Equal loading and transfer were verified by Coomassie blue staining of the membrane.

Rcho-1 trophoblast cell culture. Rcho-1 trophoblast cells were a kind gift from M. Soares. They were routinely maintained in a subconfluent condition with an RPMI 1640 culture medium supplemented with 10% fetal bovine serum (FBS), 50 µM ß-mercaptoethanol, 1 mM sodium pyruvate, 2 mM glutamine, 100 U/ml of penicillin, and 100 µg/ml of streptomycin in a humidified incubator under 5% CO₂-95% air at 37°C. Differentiation was induced by growing the cells to confluence and subsequently replacing the 10% FBS supplementation with 1% horse serum. The change to differentiation medium is indicated as day 0.

Design and cloning of lentiviral siRNA vectors. (i) Vector construction. The target sequence (19 nucleotides) was chosen in the mouse PPARß sequence 5'-GCACATCTACAACGCCTAC-3'. This sequence has identity in 18 of 19 residues with the rat PPARß sequence (the underlined residue in rat PPARß is a T) and was also efficient in the Rcho-1 cells of rat origin. A BLAST search ensured that the sequences would not target other RNAs, including PPAR mRNAs, in a nonspecific manner. The short interfering RNA (siRNA) was cloned in a lentivirus vector in which the mouse PPARß siRNA was under the control of the polymerase III-dependent H1 promoter. In addition, an internal cassette allowed the expression of the green fluorescent protein (GFP) marker gene under the control of the EF-1 promoter. A full description of the vector is given in by Wiznerowicz and Trono (65). In our study, the control vector, which contains all the features but not the siRNA, was called LV-TH and the siRNA-containing vector was named LV-THsiPPARß.

(ii) Lentivirus production. All recombinant lentiviruses were produced by transient transfection of 293T cells according to standard protocols. Briefly, subconfluent 293T cells were cotransfected with 20 µg of the control vector pLV-TH or the PPARß-targeted vector pLV-THsiPPARß, 15 µg of pCMV-R8.91, and 5 µg of pMD2G-VSVG (where CMV is cytomegalovirus and VSVG is vesicular stomatitis virus protein G) by calcium phosphate precipitation. The medium was changed after 16 h, and recombinant lentiviruses were harvested 24 h later.

(iii) Controlling lentivirus infection and silencing efficiency. The efficiency of the transduction was given by the percentage of GFP-expressing cells. At a multiplicity of infection of 60, 90% of the Rcho-1 cells expressed GFP. The efficiency of the siRNA on rat PPARß sequence was measured by transducing Rcho-1 cells with LV-THsiPPARß or the control LV-TH vector at a multiplicity of infection of 60. Forty-eight hours later, the cells were harvested, and total RNA was extracted. The level of expression of PPARß was measured by RPA.

Oil red O staining. Cultured Rcho-1 cells or sagittal sections of mouse placenta were washed in PBS and fixed in 4% paraformaldehyde for 10 min at 4°C. A fresh working solution of oil red O (Sigma Chemical, St. Louis, MO) was prepared by dilution of the oil red O stock solution (5 g/liter in 98% isopropanol) in distilled water at a ratio of 3:2. The working solution was allowed to stand for 10 min after mixing and was filtered with a 0.45-µm-pore-size filter. Subsequently, sections were stained in oil red O for 10 min, washed in tap water, and counterstained with a 0.1% methyl-green aqueous solution for 5 min. The slides were allowed to dry and were mounted with Vectashield mounting medium (Vector Laboratories, Burlingame, CA).

	RESULTS

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Growth and lethality of PPARß^–/– embryos. To assess the physiological role of PPARß in the whole organism, particularly during development, the PPARß gene was disrupted in ES cells by homologous recombination, following the strategy described in Materials and Methods and in the legend of Fig. 1. The PPARß^+/– mice (of mixed background C57BL6/Sv129) were healthy, fertile, and displayed no obvious aberrant phenotype. However, heterozygote mating did not produce the expected Mendelian ratio of mutant animals (Table 1), as the PPARß^–/– live animals were very rare (1.6% instead of the expected 25%) while no perinatal death was observed. The sacrifice of gestating mice from heterozygote mating revealed segregation ratios that were close to normal at E9.5 for the disrupted PPARß alleles among embryos. Beyond this stage, the number of live PPARß^–/– embryos decreased, and by E14.5, no living PPARß^–/– embryos were detected in the litters sacrificed, indicating that the homozygous disruption of PPARß resulted in a highly penetrant embryonic lethality. The E9.5 live PPARß^–/– mutant embryos presented various degrees of developmental retardation, with severe growth retardation but in most cases no gross abnormalities (Fig. 2A). Significantly, the placenta of PPARß^–/– concepti appeared small and abnormal, as previously observed by Barak et al. (2).

View larger version (77K): [in this window] [in a new window]   FIG. 2. Defects in extra-embryonic tissues associated with the PPARß-null genotype. (A) Morphology of E9.5 and E10.5 embryos from wt (a and c) and PPARß–/– (b and d). (B) Hematoxylin-eosin-stained sections of wt (a to c and g to i) and PPARß-null (d to f and j to l) placentas. Higher magnification views of PPARß-null placenta at E9.5 (e and f) and E10.5 (k and l) show a severe reduction of the giant cell layer in the mutant. A, heart atrium; gi, trophoblast giant cells; L, limb; la, labyrinthine trophoblast; ma, maternal decidual tissue; V, heart ventricle; sp, spongiotrophoblast. Bars = 1 mm (A), 400 µm (B, frames a, d, g, and j), and 100 µm (B, frames b, c, e, f, h, i, k, and l).

This reproducible phenotype pointed to a major role of PPARß in regulating some cellular and tissular processes taking place during placental development and was thus subjected to an in-depth analysis.

Defining the placenta alterations in PPARß^–/– concepti. For histological inspection, only placentas obtained from fetuses that were not dead in utero, as judged by a beating heart and absence of necrosis signs, were analyzed in order to exclude artifacts from postmortem placental alterations. At E9.5, the placentas of PPARß^–/– embryos, compared to those of PPARß^+/+ littermate embryos, were reduced in size and very compact (Fig. 2B). The three main placenta layers, i.e., the trophoblast giant cells, the spongiotrophoblast, and the labyrinthine layers were altered. Similar defects were observed at E10.5, showing that the defect was not merely a delay in placental development (Fig. 2B). We also sacrificed homozygous females mated with homozygous males at day 9.5 post coitum. At that stage, the litter size was close to normal (between 7 and 10 embryos), but many concepti exhibited similar placental defects as those seen in PPARß^–/– concepti from heterozygous mating (see supplemental information at http://www.unil.ch/webdav/site/cig/shared/desvergne/Nadra_et_al_supp_data.pdf). The normal litter size at E9.5, but reduced litter size at birth, indicated that lethality also occurred in homozygous matings, as expected from histological studies of the placenta. These observations established that the defect was also present in homozygous breeding, but the penetrance was somehow milder, allowing the birth of approximately 30% of the PPARß^–/– pups. Thus, these histological observations indicated that in PPARß^–/– placentas, the overall structure of the placenta is affected. Further analyses were therefore required in order to define the primary defect.

We first analyzed whether there was a spatial specificity of PPARß expression in the E9.5 developing placentas. However, in situ hybridization in wt placentas showed that PPARß mRNA is ubiquitously expressed throughout the placenta including the labyrinth, the spongiotrophoblast, and the giant cells (Fig. 3A). We thus evaluated the different trophoblast layers in the mutant placentas using a series of specific markers. The bHLH transcription factor Mash-2 is mainly expressed in the trophoblast cells of the spongiotrophoblast and labyrinth layer (23). In situ hybridization performed with a murine Mash-2 probe showed that the labyrinthine layer expressing Mash-2 was thinner in PPARß mutants (Fig. 3B). In parallel, the spongiotrophoblast cell layer, characterized by the expression of the trophoblast-specific protein Tpbpa (also called 4311) (40), was also reduced in PPARß mutant placentas, consistent with the histological analysis described above (Fig. 3B) (see also supplemental information at http://www.unil.ch/webdav/site/cig/shared/desvergne/Nadra_et_al_supp_data.pdf). The expression profile of PL-I is especially useful in characterizing early molecular events underlying trophoblast differentiation, since transcription of this gene occurs exclusively in giant cells and begins at the time of implantation (40, 47). Interestingly, we consistently observed in PPARß mutant placenta the most severe reduction in the thickness of the PL-I-expressing layer (Fig. 3B). Thus, while all layers were affected, the marker analyses underscore the severe alteration of the giant cell layer.

View larger version (65K): [in this window] [in a new window]   FIG. 3. Expression of trophoblast markers in wt and in PPARß mutant placentas. (A) Spatial expression of PPARß mRNA in E9.5 wt placenta. Left, hematoxylin-eosin (HE) staining; middle, in situ hybridization with digoxigenin-labeled antisense riboprobe for PPARß; right, control with sense riboprobe. Scale bar, 200 µm. (B) Representative E9.5 placenta sections from one wt (a, d, g, and j), and two PPARß mutant KO1 (b, e, h, and k) and KO2 (c, f, i, and l) littermates. HE corresponds to standard staining with hematoxylin-eosin. Mash-2 is a marker for spongiotrophoblast and labyrinthine trophoblast cells; Tpbpa is a specific marker of spongiotrophoblast cells; PL-I specifically labels trophoblast giant cells. Scale bars, 200 µm (a to i) and 400 µm (j to l). (C) Relative mRNA expression of PPARß, PL-I, and Id-2 in mouse placental tissues from E8.5, E9.5, E10.5, and E16.5. L27 ribosomal protein mRNA was used for normalization. Three independent samples are shown per condition. Numbers represent the relative increase (n-fold) with respect to the basal level evaluated at E8.5 time points. gi, trophoblast giant cells; la, labyrinthine trophoblast; ma, maternal decidual tissue; sp, spongiotrophoblast.

PPARß is required for trophoblastic giant cell differentiation. The putative direct role of PPARß in affecting giant cell differentiation was explored in the Rcho-1 trophoblast cell line, which provides a reliable system for the molecular analysis of the giant cell differentiation process (15, 25). When cultured in a regular 10% FBS-containing medium, these cells remain in proliferation and can be regularly subcultured while maintaining an undifferentiated status. The change from 10% FBS to 1% horse serum causes the cells to cease their proliferation and corresponds to the initiation of their differentiation into trophoblast giant cells (time zero). To evaluate the role, if any, of PPARß in this process, we exposed the cells from time zero to the PPARß agonist L-165041 and used cell morphology and PL-I expression as readouts of the differentiation status. At day 4 of the experiment, only a few control cells exhibited the features of differentiated cells; this was expected since the full differentiation process usually requires 6 to 8 days (25). Strikingly, the addition into the medium from day 0 onwards of the specific PPARß agonist L-165041 triggered a massive early differentiation (Fig. 4A) accompanied by a major increase of PL-I mRNA levels and severe reduction of Id-2 mRNA (Fig. 4B). Western blot analyses confirmed the very high levels of PL-I expressed in these cells (Fig. 4C). Because Rcho-1 cells expressed both PPARß and PPAR (Fig. 5A, top lines), the specific PPAR ligand rosiglitazone (RSG) was also tested but was found to have no effect on PL-I expression and giant cell differentiation (Fig. 4C).

View larger version (79K): [in this window] [in a new window]   FIG. 4. PPARß-dependent differentiation of Rcho-1 cells in trophoblast giant cells. Rcho-1 cells were grown to 60% confluence. The change to 1% horse serum-containing medium defines the time zero of the differentiation process. (A) Expression of PL-I in Rcho-1 cells after 4 days of differentiation in the presence of dimethyl sulfoxide (DMSO) (0.05%) used as a vehicle, the PPARß agonist L-165041 (5 µM), the PPARß agonist and the PI3K inhibitor LY294002 (10 µM). Scale bar, 80 µm. (B) Temporal expression of PL-I and Id-2 in differentiating Rcho-1 cells cultured in the presence of L-165041 or of its vehicle dimethyl sulfoxide (DMSO), as indicated. Numbers represent relative increase (n-fold) with respect to basal level in the presence of dimethyl sulfoxide (0.05%) at time zero. (C) Effect of RSG (1 µM) on PL-I expression after 4 days of treatment. Two independent duplicates per condition are shown. (D) Silencing of PPARß in Rcho-1 cells. The Rcho-1 cells were noninfected or transduced with a control lentivirus vector (LV-TH), or with a vector producing a PPARß-specific siRNA (LV-THsiPPARß). Silencing efficiency was established by measuring the mRNA levels of PPARß after 15 days of the differentiation protocol. L27 ribosomal protein mRNA was used as an internal control. (E) Differentiation of Rcho-1 cells either noninfected, transduced with LV-TH, or transduced with LV-THsiPPARß. In situ hybridization with PL-I probe was used as a marker of giant cell differentiation at day 4 of differentiation in the presence of L-165041 (5 µM) or its vehicle (dimethyl sulfoxide). Red arrowheads show some of the differentiated cells that express PL-I. The nuclei were stained with a methyl-green solution. (F) Bright-field images of Rcho-1 cells in culture after 15 days of differentiation. Ni, noninfected; Ci, LV-TH; Siß, LV-THsiPPARß/.

View larger version (61K): [in this window] [in a new window]   FIG. 5. PPARß promoting giant cell differentiation requires an intact PI3K/Akt pathway. (A) mRNA expression levels of PPARß, PPAR, PL-I, Id-2, Hand1 and I-mfa after 4 days of indicated treatments with L-165041 (5 µM) and/or PI3K inhibitor (LY294002) (10 µM). Two independent duplicates per condition are shown. (B) Western blot analysis of PL-I production in Rcho-1 cells treated by PPARß ligand and/or by LY294002. Two independent duplicates per condition are shown. The number of cells seeded in each plate was accurately quantified at the beginning of the experiment, and all culture conditions were strictly comparable. (C) mRNA expression levels of PPARß, PL-I, and I-mfa in Rcho-1 cells either noninfected (Ni), transduced with the LV-TH (Ci), or transduced with the LV-THsiPPARß (Siß), after 4 days of differentiation in the presence of either dimethyl sulfoxide, used as a vehicle, or 5 µM PPARß ligand. L27 ribosomal protein mRNA was used as an internal control. (D) Relative expression of phosphorylated Akt from Rcho-I trophoblast cells treated for 3 h with either vehicle or 5 µM PPARß ligand. Total cellular proteins from Rcho-1 cells were used for Western blot analysis. Two independent duplicates per condition are shown. (E) Relative expression of PDK1, ILK, and PTEN in Rcho-1 cells either noninfected, transduced with the LV-TH, or transduced with the LV-THsiPPARß, after 4 days of differentiation in the presence of either dimethyl sulfoxide, used as a vehicle, or 5 µM PPARß ligand. Tubulin was used as an internal control. The apparent molecular mass is indicated for each protein. Numbers represent relative increase (n-fold) with respect to the basal level in the presence of dimethyl sulfoxide (0.05%) (A and B), to basal level to (Ni) in the presence of dimethyl sulfoxide (0.05%) (C and E) or to the level at time zero in the presence of dimethyl sulfoxide (0.05%) (D). DMSO, dimethyl sulfoxide; Ni, noninfected; Ci, LV-TH; Siß, LV-THsiPPARß.

PPARß acts on giant cell differentiation via the PI3K/Akt signaling pathway and I-mfa expression. Among the factors known to contribute to giant cell differentiation, Hand1 is a bHLH transcription factor that promotes giant cell differentiation in early postimplantation mouse embryos (53, 56). In contrast, Mash-2, which is mainly expressed in the spongiotrophoblast and labyrinthine layers, as mentioned above, must be inhibited for the differentiation to occur. This is in part performed by I-mfa, a non-HLH inhibitor of the myogenic bHLH subfamily, which inhibits the transcriptional activity of Mash-2 by preventing its nuclear localization (7, 36). Id-2 acts as a negative regulator of bHLH transcription factors, and its down-regulation is required for trophoblast development (8). According to these elements, we first hypothesized that PPARß might promote giant cell differentiation via induction of Hand1 activity. However, the expression of Hand1 did not change upon PPARß-mediated induction of Rcho-1 differentiation into giant cells (Fig. 5A). In contrast, the alternative mechanism, via inhibition of Mash-2 activity, is supported by the diminished expression of Id-2 and an increased expression of I-mfa that parallels PL-I induction (Fig. 5A).

Using the same Rcho-1 cellular model, Kamei et al. (33) have described the role of the PI3K signaling pathway in giant cell differentiation. To examine whether the PPARß-mediated regulation of the different factors was dependent on the PI3K pathway, Rcho-1 cells were treated with the PPARß ligand in the presence of LY294002, a specific PI3K inhibitor. Remarkably, the addition of LY294002 efficiently prevented the PPARß-mediated up-regulation of PL-I mRNAs (Fig. 5A) and protein (Fig. 5B). This reduction of PPARß activity was also observed with respect to Id-2 repression (Fig. 5A). In contrast, the PI3K inhibitor did not affect the PPARß-induced expression of I-mfa. However, I-mfa expression remains PPARß dependent since it was abolished in PPARß siRNA-expressing cells (Fig. 5C). Further downstream of PI3K activation is the phosphorylation of Akt at the residues Ser473 and Thr308. Consistent with the results described above, treatment of Rcho-I cells with the PPARß ligand for 3 h resulted in the phosphorylation of Akt1 (Fig. 5D).

These observations indicate that PPARß activates Akt-1, which subsequently triggers the giant cell differentiation. We previously identified two kinases, namely, PDK1 and ILK, which in wounded skin regulate the PI3K/Akt pathway and which are direct PPARß target genes (11, 12). As shown in Fig. 5E, both kinases were well expressed in control Rcho-1 cells treated with the PPARß agonist, whereas they were markedly reduced in LV-THsiPPARß. We also measured the levels of phosphatase and tensin homolog deleted on chromosome 10 (PTEN), which negatively regulates the PI3K-mediated pathway by increasing the conversion from PiP3 to PiP2. Conversely, the loss of PTEN leads to Akt activation (21). PTEN levels were diminished in control Rcho-1 cells exposed to PPARß agonist but remained similar to those of the nondifferentiated cells in PPARß siRNA-treated cells (Fig. 5E).

These results demonstrate that PPARß is essential for the full differentiation of trophoblast giant cells through the activation of the PI3/Akt1 signaling pathway, in addition to a PI3K-independent induction of I-mfa.

The pathways responsible for giant cell differentiation are altered in PPARß mutant placenta. To demonstrate the relevance of the observations obtained in the Rcho-1 culture system, we biochemically characterized the trophoblastic giant cell layer by quantifying the expression levels of PL-I, Hand1, I-mfa, and Id-2 at E9.5 in control and PPARß mutant placentas. Consistent with the results obtained in Rcho-1 cells, PL-I, Hand1, and I-mfa mRNA levels were markedly reduced in PPARß mutant placenta compared with wt placenta. These decreased PL-I, Hand1, and I-mfa expressions were associated with a moderate increase in Id-2 mRNA levels (Fig. 6A).

View larger version (96K): [in this window] [in a new window]   FIG. 6. Reduced Akt activity in PPARß–/– embryos. (A) Expression profiles of PL-I, Hand1, I-mfa, and Id-2 at E9.5 in PPAR wt and mutants. L27 ribosomal protein mRNA was used as an internal control. Three independent samples are shown per genotype. (B) Relative expression of PDK1, Akt phosphorylated in Thr308 or in Ser473, and total Akt in PPARß wt and mutant placenta at E9.5. Tubulin was used as an internal control. Total cellular proteins from E9.5 placentas were used for these Western blot analyses. The apparent molecular mass is indicated for each protein. Two independent samples are shown per genotype. (C) wt E9.5 placenta. Hematoxylin-eosin staining is shown in frame a. Immunohistochemistry (b and c) reveals the Thr308-phosphorylated Akt (Akt-Thr308-P) in the nuclei of giant cells (red arrowheads). The red square in frame b indicates the region seen in frame c at higher magnification. Numbers in panels A and B represent the relative increase (n-fold) with respect to basal level to wt.

Together, these results demonstrate that the major role played by PPARß in giant cell trophoblast differentiation through activation of the PI3K/Akt1 pathway, which we characterized in cell culture, also occurs in vivo.

PPARß, lipid metabolism, and ADRP expression in giant cells. Placental fatty acid transfer is critical for normal fetal development, particularly for membrane biosynthesis, energy needs and storage, and synthesis of precursors of signaling molecules. In that respect, the labyrinth zone exerts a crucial role in regulating the bidirectional exchange between maternal and fetal compartments, with a facilitated directional flux of fatty acids from the mother to the fetus (28). Insufficient fatty acid supply may indeed result in retarded fetal growth (9). Little is yet known about the molecular mechanism of fatty acid transfer, while even less is known about fatty acid metabolism in the placenta in the early stages, when the labyrinth is not yet fully developed. The severe growth retardation observed in the PPARß mutant embryos at E9.5 (Fig. 2) and the role of PPARß in lipid metabolism (3, 17) led us to investigate the lipid status in E9.5 placentas, in comparison with the situation observed at E16.5.

As a first step, we assessed the presence and location of lipid droplets in the placenta, by oil red O staining of tissue sections of placentas. Strikingly, lipid accumulation was already observed at E9.5, remarkably pronounced in the giant cells (Fig. 7A, frames a to d) and to a lesser extent in the labyrinth part. At E16.5, the giant cells were no longer distinguishable, and lipid droplets were abundantly found in both the maternal decidua and the labyrinth, while they were undetectable in the spongiotrophoblast (Fig. 7A, frames e to h). This lipid accumulation in the form of cytosolic droplets was paralleled by the expression pattern of ADRP, a major lipid droplet-associated protein that helps in packaging neutral lipids into discrete lipid storage droplets in the cytoplasm (31). At E9.5, ADRP immunoreactivity was mainly localized in the cytoplasm of trophoblast giant cells (Fig. 7B, frames a to d). Higher magnification revealed the particular pattern of ADRP protein localized as brown rings around the lipid droplets (Fig. 7B, frame d). In the labyrinthine trophoblast cells, ADRP was found toward the apical surface of the cells and was concentrated along the fetal vessels (Fig. 7B). At E16.5, ADRP was abundantly expressed in the decidua and the labyrinth (Fig. 7B, frames e to h), closely reproducing the lipid droplet pattern.

View larger version (127K): [in this window] [in a new window]   FIG. 7. Lipid storage droplets accumulate in the mouse placenta. (A) Oil red O staining of sagittal sections of mouse placenta at E9.5 (a to d) and E16.5 (e to h). Abundant small lipid droplets were observed inside the cytoplasm of trophoblast giant cells. As seen in frame d, the droplets are particularly localized near the nucleus. Black arrowheads indicate some lipid droplets. Bars: 200 µm (a and e), 40 µm (b, c, and f), 20 µm (g and h), and 8 µm (d). (B) Distribution of ADRP in mouse placenta at E9.5 (a to d) and E16.5 (g to j). Red arrowheads indicate some specific labeling corresponding to ADRP protein. (C) Expression profile of ADRP mRNA in E9.5 placentas from PPARß wt and mutant concepti (left) and from PPAR wt and mutant concepti (right). (D) Relative mRNA expression profiles of ADRP and PPAR in mouse placental tissues from E8.5, E9.5, E10.5, and E16.5. L27 ribosomal protein mRNA was used for normalization. Numbers represent the relative increase (n-fold) with respect to the basal level at E8.5 (D) or to wt (C). gi, trophoblast giant cells; la, labyrinthine trophoblast; ma, maternal decidual tissue; sp, spongiotrophoblast.

ADRP expression is associated to giant cell differentiation and under the control of PPARß. To evaluate whether ADRP expression and lipid droplet accumulation in the giant cells were due to the placental environment or could be precisely attributed to giant cell functions, we furthered our investigations in Rcho-1 cells. Amazingly, an increase in ADRP expression and the appearance of lipid droplets accompanied giant cell differentiation, obtained either using the classical procedure or by accelerated differentiation via exposure to a PPARß ligand. In this context, PPAR ligand RSG provoked little if any increased ADRP expression. The need for PPARß was further shown in cells infected by LV-THsiPPARß, which prevented the PPARß ligand-induced expression of ADRP (Fig. 8A).

View larger version (69K): [in this window] [in a new window]   FIG. 8. ADRP expression is associated to giant cell differentiation. (A) Expression profile of ADRP in Rcho-1 cells after 4 days of treatment with L-165041 (5 µM), RSG (1 µM), or PI3K inhibitor (LY294002) (10 µM) (left and bottom panels) and in cells either noninfected (Ni), transduced with the LV-TH (Ci), or transduced with the LV-THsiPPARß (Siß) (right panel). Dimethyl sulfoxide (DMSO) was used as a vehicle. (B) Relative mRNA expression profiles of ADRP and aP2 in mouse placental tissues at E16.5, in white adipose tissue (WAT), and in heart. L27 ribosomal protein mRNA was used for normalization. Numbers represent the relative increase (n-fold) with respect to basal level in the presence of dimethyl sulfoxide (0.05%) (A) or to level in white adipose tissue (B).

In summary, our observations demonstrate the crucial role of PPARß in the differentiation of the trophoblastic giant cell layer via two overlapping molecular mechanisms which involve a PPARß-mediated increased Akt1 activity and a direct positive regulation of the transcription factor I-mfa. We further demonstrated that giant cells accumulate lipid, together with the expression of the ADRP protein, conferring on the giant cells an unexpected role in lipid metabolism.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

 
The analysis of PPARß functions in the mouse placenta provides important information. First, although PPARß and PPAR are expressed in the three layers of the developing placenta, functional redundancies or compensations between them, if any, are not fully operative. Second, PPARß is a major regulator of the differentiation of the secondary giant cells, which play a critical role in the establishment of the placental structure and fulfill an important endocrine function. Third, we reveal that secondary giant cells are the prime site of lipid accumulation in the developing placenta, an event which is also under the control of PPARß.

PPARß and embryonic lethality. Conflicting results concerning the embryonic lethality of PPARß null mutant mice appeared in two previous reports (51, 2). Indeed, Peters et al. (51) observed no embryonic lethality on a pure C57BL/6N background but a lower than expected number of newborn homozygous pups on a mixed genetic background. In contrast, Barak et al. (2) reported a severe embryonic lethality in either Sv129 or C57BL/6J background. Our observations are closer to those of Barak et al. (2), with a frequent embryonic lethality between E9.5 and E10.5 due to an altered placenta formation. An explanation of these discrepancies may lie in the mutation performed on the PPARß gene, with a disruption of the DNA-binding domain therein, as in Barak et al. (2), whereas the mutation performed by Peters et al. (51) disrupted the last 60 amino acids of the ligand-binding domain. The latter approach might have resulted in a hypomorph allele, retaining some aporeceptor functions. A comparative analysis of these two models of PPARß disruption in a congenic background might reveal very interesting features of PPARß functions.

The fact that we were able to obtain some productive homozygous pairs, in which the defects in the placenta were still present but less penetrant, is likely due to the presence of one or more modifiers responsible for the higher survival rate of PPARß^–/– embryos. However, while we have not been able so far to improve the survival rate by enriching the mixed genetic background toward Sv129 or toward C57BL/6 background, a contribution of the genetic background to the observed phenotypes cannot be excluded. Large-scale and in-depth genetic studies will be needed to identify the genes that are partners in this complex trait.

PPARß promotes giant cell differentiation. We demonstrate that deletion of PPARß leads to a dramatic decrease of the trophoblast secondary giant cell layer in vivo and abolishes the ability of Rcho-1 cells to differentiate into giant cells in cell culture. Conversely, agonist activation of PPARß in a cell culture model of trophoblast giant cell differentiation markedly accelerates and increases the extent of differentiation.

Our studies contribute to identifying the factors involved in the development and endocrine functions of the placenta. Members of the bHLH factor family are involved in the control of commitment, differentiation, and development of many different tissues and organs, including the placenta (23). Hand1 promotes the differentiation of trophoblast giant cells (53). In contrast, Mash-2 has the opposite effect as it is required to maintain the pool of precursors in the ectoplacental cone and spongiotrophoblast, and its sustained expression precludes their differentiation in giant cells (23). Id-2 belongs to a subset of HLH proteins and acts as a dominant negative factor by dimerizing and sequestering the heterodimerization partner of bHLH transcription factors (48). In Rcho-1 cells, overexpression of a related protein (Id-1) inhibits their differentiation into giant cells (8). This can be paralleled with the expression of Id-2, which is undetectable in differentiated trophoblast giant cells but remains expressed in the extra-embryonic ectoderm of the chorion (30). Finally, the bHLH-repressor protein I-mfa is also required for the generation of trophoblast giant cells, possibly acting as a direct inhibitor of the activity of Mash-2 by preventing its nuclear import (36). In the present study, we demonstrate that PPARß activation does not affect Hand1 expression but increases I-mfa mRNA levels in a PI3K-independent manner and decreases Id-2 expression in a PI3K-dependent manner. As proposed in Fig. 9 and its legend, these two likely independent mechanisms are converging in trophoblast cell differentiation. Interestingly, the I-mfa mutants exhibit a generally similar phenotype to that of PPARß mutant embryos, with an overall reduced size of the embryos, a lack of trophoblast giant cells, and a lethality that occurs at mid-gestation (36).

View larger version (11K): [in this window] [in a new window]   FIG. 9. PPARß plays a key role in trophoblast giant cell differentiation via two converging mechanisms. Hand1 and Mash-2 are two crucial transcription factors involved in promoting and inhibiting giant cell differentiation, respectively. In addition, Id-2 acts as a dominant negative factor by dimerizing and sequestering the heterodimerization partner of bHLH transcription factors and must be down-regulated to allow trophoblast giant cell differentiation. PPARß first acts by increasing Akt activity, which leads to Id-2 down-regulation. The bHLH transcription factor targeted by Id-2 in the trophoblast cells is not ascertained but could include Hand1 (dotted line) or the heterodimerization partner of Hand1, which in the giant cells is not yet identified (56). The role of PPARß in directly regulating Hand1 is unclear since Hand1 expression is not modified in the Rcho-1 cell model. Secondly, PPARß also increases the expression of I-mfa, which acts as an inhibitor of Mash-2 activity, possibly by impairing its nuclear import. Finally, the subsequent differentiation is associated with ADRP expression and lipid accumulation, which also depends on PPARß and its activity on Akt-1.

PPARß and lipid metabolism in the placenta. Our present results strongly support two mechanisms, possibly independent, of lipid storage and ADRP expression in the placenta. At an early stage, when giant cells are forming the main interface between the maternal and the fetal compartments, giant cells are the major site of lipid accumulation, which is accompanied by increased ADRP expression. Herein, we demonstrate that this early ADRP increased expression is PPARß dependent. At a later stage, upon major development of a functional labyrinth structure, the labyrinth and the maternal decidua become the main sites where lipid droplets are observed, and a careful examination points to the major presence of lipids in the cells that surround the fetal vessels. This labyrinth expression is possibly dependent on PPAR, as previously suggested (1, 55).

These features are particularly remarkable since both PPARß and PPAR are expressed throughout the three placenta layers, suggesting that the specificity of their action must depend on distinct activation processes, including ligand availability and/or target gene specificity. In mice and in humans, ADRP is a known PPAR target gene, and a PPRE identified in the ADRP promoter mediates its response to the PPARß agonist GW501516 (6, 62) and also to PPAR and to PPAR (24, 14, 57). In Rcho-1 cells, the full activity of PPARß to increase the expression of ADRP requires an intact PI3K/Akt1 signaling pathway (Fig. 8) and a successful differentiation in giant cells. Thus, the high level of ADRP mRNA in Rcho-1 differentiated cells is likely the result of two processes: direct transcriptional activity of PPARß at the ADRP promoter and indirect action of PPARß on differentiation via the PI3K/Akt1 pathway.

Although elucidation of its function is still in progress, ADRP seems to be involved in the formation and stabilization of lipid droplets (19). Recombinant expression of ADRP in COS-7 cells demonstrated that this protein facilitates the transport of long-chain free fatty acids and may function as a fatty acid carrier protein (18). Our results raise the question as to whether or not lipid accumulation in the giant cells during the early stages of placental development fulfills the same role as it does in the labyrinth cells at a later stage. In the latter, the ADRP location in close proximity to the vessel walls suggests a metabolic role in feto-maternal exchange. This nutritional role in the giant cells cannot be excluded. However, ADRP not only encircles droplets mainly composed of triglycerides, such as those found in the lactating mammary gland, but also droplets rich in cholesterol stored as steroid hormone precursors in adrenocortical cells (27). In the rodent placentas, androgen synthesis takes places within trophoblast giant cells (32). Rcho-1 cells synthesize both progesterone and androstenedione (66). Thus, lipid droplet accumulation and ADRP expression in trophoblast giant cells may rather contribute to the production of steroid hormones. Therefore, PPARß null concepti would suffer from both a low number of differentiated giant cells and an additional alteration of their endocrine functions via decreased levels of ADRP, consequently impairing placenta and embryonic growth.

In conclusion, we have revealed some crucial regulatory events in giant cell differentiation mediated by PPARß. These results also bring to light the significant role of PPARß in correlation with lipid accumulation, which may fulfill an endocrine function as well as contribute to nutrient exchange between the fetus and the mother. Collectively, these observations concerning PPARß null placentas emphasize the need to carefully evaluate the effects of PPAR ligands on placenta development and physiology when they are considered for use as therapeutic drugs in chronic metabolic diseases, which may also affect pregnant women.

	ACKNOWLEDGMENTS

 
We are grateful to D. Duboule and J. Zakany for help in obtaining the PPARß mutant mice as well as for stimulating discussion. We also thank D. Metzger and P. Chambon for comments on the manuscript and M. J. Soares for making the Rcho-I cell line available to us. We thank M. Mark for helpful advice, J. Rossant for generous gifts of PL-I and Tpbpa probes, F. Guillemot for providing us with the Mash-2 cDNA, and C. Schneider and D. Galaud for technical assistance.

This work was supported by grants from Swiss National Science Foundation to B.D. and W.W.

	FOOTNOTES

 
* Corresponding author. Mailing address: Center for Integrative Genomics, University of Lausanne, CH-1015 Lausanne, Switzerland. Phone: 41 21 6924140. Fax: 41 21 6924115. E-mail: beatrice.desvergne{at}unil.ch.

Present address: School of Biological Sciences, Nanyang Technological University, 60 Nanyang Drive, Singapore 637551, Singapore.

Present address: Department of Zoology, University of Dehli, Dehli 110007, India.

http://www.frontiers-in-genetics.org/en/index.php.

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