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The Organization, Promoter Analysis, and Expression of the Human PPARγ Gene*

Open AccessPublished:July 25, 1997DOI:https://doi.org/10.1074/jbc.272.30.18779
      PPARγ is a member of the PPAR subfamily of nuclear receptors. In this work, the structure of the human PPARγ cDNA and gene was determined, and its promoters and tissue-specific expression were functionally characterized. Similar to the mouse, two PPAR isoforms, PPARγ1 and PPARγ2, were detected in man. The relative expression of human PPARγ was studied by a newly developed and sensitive reverse transcriptase-competitive polymerase chain reaction method, which allowed us to distinguish between PPARγ1 and γ2 mRNA. In all tissues analyzed, PPARγ2 was much less abundant than PPARγ1. Adipose tissue and large intestine have the highest levels of PPARγ mRNA; kidney, liver, and small intestine have intermediate levels; whereas PPARγ is barely detectable in muscle. This high level expression of PPARγ in colon warrants further study in view of the well established role of fatty acid and arachidonic acid derivatives in colonic disease. Similarly as mouse PPARγs, the human PPARγs are activated by thiazolidinediones and prostaglandin J and bind with high affinity to a PPRE. The human PPARγ gene has nine exons and extends over more than 100 kilobases of genomic DNA. Alternate transcription start sites and alternate splicing generate the PPARγ1 and PPARγ2 mRNAs, which differ at their 5′-ends. PPARγ1 is encoded by eight exons, and PPARγ2 is encoded by seven exons. The 5′-untranslated sequence of PPARγ1 is comprised of exons A1 and A2, whereas that of PPARγ2 plus the additional PPARγ2-specific N-terminal amino acids are encoded by exon B, located between exons A2 and A1. The remaining six exons, termed 1 to 6, are common to the PPARγ1 and γ2. Knowledge of the gene structure will allow screening for PPARγ mutations in humans with metabolic disorders, whereas knowledge of its expression pattern and factors regulating its expression could be of major importance in understanding its biology.
      White adipose tissue is composed of adipocytes, which play a central role in lipid homeostasis and the maintenance of energy balance in vertebrates. These cells store energy in the form of triglycerides during periods of nutritional affluence and release it in the form of free fatty acids at times of nutritional deprivation. Excess of white adipose tissue leads to obesity (
      • Auwerx J.
      • Martin G.
      • Guerre-Millo G.
      • Staels B.
      ,
      • Flier J.S.
      ,
      • Spiegelman B.M.
      • Flier J.S.
      ), whereas its absence is associated with lipodystrophic syndromes (
      • Moller D.E.
      • Flier J.S.
      ). In contrast to the development of brown adipose tissue, which mainly takes place before birth, the development of white adipose tissue is the result of a continuous differentiation/development process throughout life (
      • Flier J.S.
      ,

      Auwerx, J., Schoonjans, K., Fruchart, J. C., and Staels, B. (1996)Atherosclerosis , 124, (suppl.) S29–S37.

      ). During development, cells that are pluripotent become increasingly restricted to specific differentiation pathways. Adipocyte differentiation results from coordinate changes in the expression of several proteins, which are mostly involved in lipid storage and metabolism, that give rise to the characteristic adipocyte phenotype. The changes in expression of these specialized proteins are mainly the result of alterations in the transcription rates of their genes.
      Several transcription factors including the nuclear receptor PPARγ (
      • Tontonoz P.
      • Hu E.
      • Graves R.A.
      • Budavari A.I.
      • Spiegelman B.M.
      ,
      • Tontonoz P.
      • Hu E.
      • Spiegelman B.M.
      ), the family of CCAATT enhancer binding proteins (C/EBP)
      The abbreviations used are: C/EBP, CCAATT enhancer binding protein; LPL, lipoprotein lipase; kb, kilobase(s); bp, base pair(s); UTR, untranslated region; RACE, rapid amplification of cDNA ends; EMSA, electrophoretic mobility shift assays; ACO, acyl-CoA oxidase; PAC, P1-derived artificial chromosome; PPRE, peroxisome proliferator response element; NIDDM, non-insulin-dependent diabetes mellitus.
      1The abbreviations used are: C/EBP, CCAATT enhancer binding protein; LPL, lipoprotein lipase; kb, kilobase(s); bp, base pair(s); UTR, untranslated region; RACE, rapid amplification of cDNA ends; EMSA, electrophoretic mobility shift assays; ACO, acyl-CoA oxidase; PAC, P1-derived artificial chromosome; PPRE, peroxisome proliferator response element; NIDDM, non-insulin-dependent diabetes mellitus.
      (
      • Freytag S.O.
      • Geddes T.J.
      ,
      • Freytag S.O.
      • Paielli D.L.
      • Gilbert J.D.
      ,
      • Christy R.J.
      • Yang V.W.
      • Ntambi J.M.
      • Geiman D.E.
      • Landschulz W.H.
      • Friedman A.D.
      • Nakabeppu Y.
      • Kelly T.J.
      • Lane M.D.
      ,
      • Wu Z.
      • Xie Y.
      • Bucher N.L.R.
      • Farmer S.R.
      ,
      • Wu Z.
      • Bucher N.L.R.
      • Farmer S.R.
      ,
      • Yeh W.C.
      • Cao Z.
      • Classon M.
      • McKnight S.
      ) and the basic helix-loop-helix leucine zipper transcription factor ADD1/SREBP1 (
      • Tontonoz P.
      • Kim J.B.
      • Graves R.A.
      • Spiegelman B.M.
      ,
      • Kim J.B.
      • Spiegelman B.M.
      ) orchestrate the adipocyte differentiation process (for reviews, see Refs.
      • Auwerx J.
      • Martin G.
      • Guerre-Millo G.
      • Staels B.
      ,
      • Spiegelman B.M.
      • Flier J.S.
      ,
      • Schoonjans K.
      • Staels B.
      • Auwerx J.
      ,
      • Schoonjans K.
      • Staels B.
      • Auwerx J.
      ,
      • Cornelius P.
      • MacDougald O.A.
      • Lane M.D.
      ). In contrast to the wide tissue distribution of the various C/EBPs, PPARγ has been shown to have an adipose-restricted pattern of expression in mouse. The currently favored hypothesis is that C/EBPβ and δ induce the expression of PPARγ (
      • Wu Z.
      • Xie Y.
      • Bucher N.L.R.
      • Farmer S.R.
      ), which then triggers the adipogenic program. Terminal differentiation then requires the concerted action of both PPARγ, C/EBPα, and ADD-1/SREBP1 (
      • Tontonoz P.
      • Hu E.
      • Spiegelman B.M.
      ,
      • Kim J.B.
      • Spiegelman B.M.
      ). Several arguments support the important role of PPARγ in adipocyte differentiation. First, overexpression of PPARγ by itself can induce adipocyte conversion of fibroblasts (
      • Tontonoz P.
      • Hu E.
      • Graves R.A.
      • Budavari A.I.
      • Spiegelman B.M.
      ). In addition, PPARγ together with C/EBPα can induce transdifferentiation of myoblasts into adipocytes (
      • Hu E.
      • Tontonoz P.
      • Spiegelman B.M.
      ). Second, the description of functional PPREs in the regulatory sequences of several of the genes that are induced during adipocyte differentiation, such as the genes coding for adipocyte fatty acid binding protein, aP2 (
      • Tontonoz P.
      • Hu E.
      • Graves R.A.
      • Budavari A.I.
      • Spiegelman B.M.
      ), phosphoenolpyruvate carboxykinase (PEPCK) (
      • Tontonoz P.
      • Hu E.
      • Devine J.
      • Beale E.G.
      • Spiegelman B.M.
      ), acyl-CoA synthetase (ACS) (
      • Schoonjans K.
      • Staels B.
      • Grimaldi P.
      • Auwerx J.
      ,
      • Schoonjans K.
      • Watanabe M.
      • Suzuki H.
      • Mahfoudi A.
      • Krey G.
      • Wahli W.
      • Grimaldi P.
      • Staels B.
      • Yamamoto T.
      • Auwerx J.
      ), and lipoprotein lipase (LPL) (
      • Schoonjans K.
      • Peinado-Onsurbe J.
      • Heyman R.
      • Briggs M.
      • Cayet D.
      • Deeb S.
      • Staels B.
      • Auwerx J.
      ), is consistent with the crucial role attributed to PPARγ in adipocyte differentiation. Finally, PPAR activators, such as fibrates (
      • Brandes R.
      • Hertz R.
      • Arad R.
      • Naishtat S.
      • Weil S.
      • Bar-Tana J.
      ,
      • Gharbi-Chibi J.
      • Teboul M.
      • Bismuth J.
      • Bonne J.
      • Torresani J.
      ) and fatty acids (
      • Tontonoz P.
      • Hu E.
      • Spiegelman B.M.
      ,
      • Amri E.-Z.
      • Bertrand B.
      • Ailhaud G.
      • Grimaldi P.
      ,
      • Chawla A.
      • Lazar M.A.
      ,
      • Kliewer S.A.
      • Lenhard J.M.
      • Willson T.M.
      • Patel I.
      • Morris D.C.
      • Lehman J.M.
      ), or synthetic PPARγ ligands, such as the thiazolidinediones (
      • Tontonoz P.
      • Hu E.
      • Spiegelman B.M.
      ,
      • Kliewer S.A.
      • Lenhard J.M.
      • Willson T.M.
      • Patel I.
      • Morris D.C.
      • Lehman J.M.
      ,
      • Forman B.M.
      • Tontonoz P.
      • Chen J.
      • Brun R.P.
      • Spiegelman B.M.
      • Evans R.M.
      ), induce adipocyte differentiation. In this context, it is interesting to note that prostanoids, which are potent inducers of adipose differentiation programs (
      • Gaillard D.
      • Negrel R.
      • Lagarde M.
      • Ailhaud G.
      ,
      • Negrel R.
      • Gaillard D.
      • Ailhaud G.
      ,
      • Aubert J.
      • Ailhaud G.
      • Negrel R.
      ), may be one of the natural ligands of PPARγ. In addition to PPARγ, PPARα, but not PPARδ, has been shown to have some, albeit weaker, adipogenic activity (
      • Brun R.P.
      • Tontonoz P.
      • Forman B.M.
      • Ellis R.
      • Chen J.
      • Evans R.M.
      • Spiegelman B.M.
      ).
      To better understand the physiological role of PPARγ in human physiology, it is crucial that we gain insight into the regulation of PPARγ gene expression in man. Therefore, we cloned the human PPARγ cDNAs, determined the structure of the human PPARγ gene, and studied the expression of the PPARγ mRNAs and the regulation of their promoter. Both PPARγ1 and 2 are produced in human tissues but PPARγ2 appears to be the minor isoform in man. In addition to adipose tissue, which contains high levels of PPARγ, we demonstrate high level expression of human PPARγ in the colon. The structure of the gene encoding the mouse and human PPARγs is highly conserved. Furthermore our results demonstrate that 3 and 1 kb of DNA upstream of the transcription start sites of PPARγ1 and γ2, respectively, are sufficient to control basal and tissue-specific PPARγ gene expression.

      EXPERIMENTAL PROCEDURES

      Materials and Oligonucleotides

      The oligonucleotides used for various experiments in this manuscript are listed in Table I.
      Figure thumbnail gr1
      Figure 1Sequence of the human PPARγ cDNA and comparison with the mouse PPARγ sequence. A, sequence comparison of mouse and human PPARγ. Identical amino acids are indicated by a vertical line and conservative changes are indicated by a dot. B, splicing of exon A2 and exon B with exon 1. hPPARγ1 contains two extra amino acids relative to mPPARγ1. The presence of a promoter in front of exon A and B is indicated by an arrow. Nucleotides incapitals are located in exons, whereas the nucleotides in the intron are in lowercase.
      Table IOligonucleotides used in this study listed from 5′ to 3′
      NameSequence
      LF-2TCTCCGGTGTCCTCGAGGCCGACCCAA
      LF-14AGTGAAGGAATCGCTTTCTGGGTCAAT
      LF-18AGCTGATCCCAAAGTTGGTGGGCCAGA
      LF-20CATTCCATTCACAAGAACAGATCCAGTGGT
      LF-21GGCTCTTCATGAGGCTTATTGTAGAGCTGA
      LF-22GCAATTGAATGTCGTGTCTGTGGAGATAA
      LF-23GTGGATCCGACAGTTAAGATCACATCTGT
      LF-24GTAGAAATAAATGTCAGTACTGTCGGTTTC
      LF-25TCGATATCACTGGAGATCTCCGCCAACAG
      LF-26ACATAAAGTCCTTCCCGCTGACCAAAGCAA
      LF-27CTCTGCTCCTGCAGGGGGGTGATGTGTTT
      LF-28GAAGTTCAATGCACTGGAATTAGATGACA
      LF-29GAGCTCCAGGGGTTGTAGCAGGTTGTCTT
      LF-33GACGGGCTGAGGAGAAGTCACACTCTGA
      LF-35AGCATGGAATAGGGGTTTGCTGTAATTC
      LF-36TAGTACAAGTCCTTGTAGATCTCC
      LF-44GTCGGCCTCGAGGACACCGGAGAG
      LF-58CACTCATGTGACAAGACCTGCTCC
      LF-59GCCGACACTAAACCACCAATATAC
      LF-60CGTTAAAGGCTGACTCTCGTTTGA
      AII J PPREGATCCTTCAACCTTTACCCTGGTAGA
      ACO PPREGATCCCGAACGTGACCTTTGTCCTGGTCCC
      LPL PPREGATCCGTCTGCCCTTTCCCCCTCTTCA
      γASGCATTATGAGCATCCCCAC
      γSTCTCTCCGTAATGGAAGACC
      γ2SGCGATTCCTTCACTGATAC
      CDSTTCTAGAATTCAGCGGCCGC(T)30(G/A/C)(G/A/C/T)

      Isolation of the Human PPARγ cDNA and Gene, Restriction Mapping, Determination of Intron/Exon Boundaries, and DNA Sequencing

      A human adipose tissue λgt11 library was screened with a random primed 32P-labeled 200 bp fragment, covering the DNA-binding domain of the mouse PPARγ cDNA. After hybridization, filters were washed in 2 × SSC, 0.1% SDS for 10 min at 20 °C and twice for 30 min in 1 × SSC, 0.1% SDS at 50 °C and subsequently exposed to x-ray film (X-OMAT-AR, Kodak). Of several positive clones, one clone 407 was characterized in detail. The insert of this clone, starting ±90 bp upstream of the ATG start codon and extending downstream into the 3′-untranslated region (UTR) sequence, was subcloned in the EcoRI site of pBluescript SK to generate clone 407.2. Sequence analysis of 407.2 confirmed it as being the human homologue of the mouse PPARγ2 cDNA. While this work was in progress, other groups also reported the isolation of human PPARγ2 cDNA clones (
      • Elbrecht A.
      • Chen Y.
      • Cullinan C.A.
      • Hayes N.
      • Leibowitz M.D.
      • Moller D.E.
      • Berger J.
      ,
      • Lambe K.G.
      • Tugwood J.D.
      ).
      To isolate genomic P1-derived artificial chromosome (PAC) clones containing the entire human PPARγ gene, the primer pair LF-3 and LF-14 was used to amplify an 86-bp probe with human genomic DNA as template. This fragment was then used to screen a PAC human genomic library from human foreskin fibroblasts. Three positive clones, P-8854, P-8855, and P-8856, were isolated. Restriction digestion and Southern blotting were performed according to classical protocols as described by Sambrook et al. (
      • Sambrook J.
      • Fritsch E.F.
      • Maniatis T.
      ). Sequencing reactions were performed, according to the manufacturer instructions, using the T7 sequencing kit (Pharmacia Biotech Inc.).

      Determination of the Transcription Initiation Site: Primer Extension and 5′-Rapid Amplification of cDNA Ends (5′-RACE)

      Primer Extension

      The oligonucleotide LF-35 was32P-labeled with T4-polynucleotide kinase (Amersham Life Science, Inc) to a specific activity of 107 dpm/50 ng and purified by gel electrophoresis. For primer extension, 105dpm of oligonucleotide was added in a final volume of 100 μl to 50 μg of adipose tissue total RNA isolated from different patients. Primer extension analysis was performed following standard protocols utilizing a mixture of 1.25 units of avian mycloblastosis virus reverse transcriptase (Life Technologies, Inc.) and 100 units of Moloney murine leukemia virus reverse transcriptase (Life Technologies, Inc.). A sequencing reaction and molecular mass standards were used to map the 5′-end of the extension products.

      5′-RACE

      The Marathon cDNA amplification kit (CLONTECH) was used to obtain a library of adaptor-ligated double-stranded cDNA from human adipose tissue. 1 μg of poly(A)+ RNA was used as a template for the first strand synthesis, with the 52-mer CDS primer and 100 units of the MMLV reverse transcriptase in a total volume of 10 μl. Synthesis was carried out at 42 °C for 1 h. Next, the second strand was synthetized at 16 °C for 90 min in a total volume of 80 μl containing the enzyme mixture (RNase H, Escherichia coli DNA polymerase I, and E. coli DNA ligase), the second strand buffer, the dNTP mix, and the first strand reaction. cDNA ends were then made blunt by adding to the reaction 10 units of T4 DNA polymerase and incubating at 16 °C for 45 min. The double-stranded cDNA was phenol/chloroform extracted, ethanol precipitated, and resuspended in 10 μl of water. Half of this volume was used to ligate the adaptor to the cDNA ends (adaptor sequence CTAATACGACTCACTATAGGGCTCGAGCGGCCGCCCGGGCAGGT) in a total volume of 10 μl using 1 unit of T4 DNA ligase. The ligation reaction was incubated 16 h at 16 °C. The resulting cDNA library was diluted to a final concentration of 0.1 mg/ml.
      The 5′-end of PPARγ1 was PCR-amplified using 5 μl of the library as a template with the oligonucleotides AP-1 (binding to the adaptor) and LF-45 (binding antisense to the 5′-end of the PPARγ1). After an initial denaturing step at 95 °C for 3 min, 25 cycles were done at the following conditions: 10 s at 95 °C, 20 s at 60 °C, and 30 s at 72 °C. The resulting PCR product was reamplified for 30 additional cycles at the same conditions using the nested oligonucleotides AP2 (nested to AP1) and LF-2 (nested to LF-45). The PCR product was analyzed on a 2% agarose gel, treated withPfu polymerase (Stratagene) and cloned into theEcoRV site of pBluescript SK+. A total of 20 white colonies were grown and sequenced from both ends using the oligonucleotides T3 and T7 (Dye Terminator Cycle sequencing kit, Applied Biosystems).
      For the determination of the 5′-end of PPARγ2, the same procedure was followed except that the oligonucleotide LF-14 (specific for the PPARγ2 5′-UTR) was used in the first round PCR, and the oligonucleotide LF-35 (nested to LF-14) was used in the second round PCR with the same cycling conditions.

      Tissue Biopsies and Cell Culture

      Omental adipose tissue, small and large intestine, kidney, muscle, and liver biopsies were obtained from non-obese adult subjects undergoing elective surgery or endoscopy. All subjects had fasted overnight before surgery (between 8.00 p.m. and 10 a.m.) and received intravenous saline infusion. They had given informed consent, and the project was approved by the ethics committee of the University of Lille. All tissue was immediately frozen in liquid nitrogen until RNA preparation.
      Standard cell culture conditions were used to maintain 3T3-L1 (obtained from ATCC), CV-1 (a kind gift from Dr. R. Evans, Salk Institute, La Jolla, CA), and Hep G2 cells (ATCC). BRL-49,653, supplied by Ligand Pharmaceuticals, San Diego, CA (in DMSO) and fatty acids (in ethanol) were added to the medium at the concentrations and times indicated. Control cells received vehicle only. Fatty acids were complexed to serum albumin contained in delipidated and charcoal-treated fetal calf serum by preincubation for 45 min at 37 °C.

      mRNA Analysis by RT-Competitive PCR Assay

      RNA preparation of total cellular RNA was performed as described previously (
      • Saladin R.
      • De Vos P.
      • Guerre-Millo M.
      • Leturque A.
      • Girard J.
      • Staels B.
      • Auwerx J.
      ). The absolute mRNA concentration of the differentially spliced PPARγ variants was measured by reverse transcription reaction followed by competitive polymerase chain reaction (RT-competitive PCR) in the presence of known amounts of competitor DNA yielding amplicons of different size allowing the separation and the quantification of the PCR products. The competitor was constructed by deletion of a 74-bp fragment (nucleotides +433 to +507 by HindIII digestion) of PPARγ1 cloned into pBluescript KS+, yielding pBSCompPPARγ. Working solution of the competitor was prepared by in vitro transcription followed by serial dilution in 10 mm Tris-HCl (pH 8.3), 1 mm EDTA buffer. For RT-competitive PCR, the antisense primer hybridized to the 3′-end of exon 3 (γAS:5′-GCATTATGAGCATCCCCAC-3′, nt +600 to +620) and the sense primer to exon 1 (γS:5′-TCTCTCCGTAATGGAAGACC-3′, nt +146 to +165) or to the B exon (γ2S:5′-GCGATTCCTTCACTGATAC-3′, nt +41 to +59). Therefore, the same competitor served to measure either total PPARγ mRNAs (γ1 + γ2; with primers γAS and γS) or, specifically, PPARγ2 mRNA (with primers γAS and γ2S). The γAS/γS primer pair gave PCR products of 474 and 400 bp for the PPARγ mRNAs and competitor, respectively. The primer pair γAS/γ2S gave 580 bp for PPARγ2 mRNA and 506 bp for the competitor. For analysis of the PCR products, the sense primers γS and γ2S were 5′-end labeled with the fluorescent dye Cy-5 (Eurogentec, Belgium).
      First-strand cDNA synthesis was performed from total RNA (0.1 μg) in the presence of the antisense primer γAS (15 pmol) and of thermostable reverse transcriptase (2.5 units; Tth DNA polymerase, Promega) as described (
      • Vidal H.
      • Auboeuf D.
      • De Vos P.
      • Staels B.
      • Riou J.P.
      • Auwerx J.
      • Laville M.
      ). After the reaction, half of the RT volume was added to the PCR mix (90 μl) containing the primer pair γAS/γS for the assay of PPARγ total mRNA, whereas the other half was added to a PCR mix (10 mm Tris-HCl, pH 8.3, 100 mm KCl, 0.75 m EGTA, 5% glycerol, 0.2 mm dNTP, 5 units of Taq polymerase) containing the primer pair γAS/γ2S for the assay of PPARγ2 mRNA. Four aliquots (20 μl) of the mixture were then transferred to microtubes containing a different, but known, amount of competitor. After 120 s at 95 °C, the samples were subjected to 40 PCR cycles (40 s at 95 °C, 50 s at 55 °C, and 50 s at 72 °C). The fluorescent-labeled PCR products were analyzed by 4% denaturing polyacrylamide gel electrophoresis using an automated laser fluorescence DNA sequencer (ALFexpress, Pharmacia, Uppsala, Sweden), and integration of the area under the curve using the Fragment manager software (Pharmacia) was performed as described (
      • Vidal H.
      • Auboeuf D.
      • De Vos P.
      • Staels B.
      • Riou J.P.
      • Auwerx J.
      • Laville M.
      ).
      To validate this technique, human PPARγ2 mRNA was synthesized byin vitro transcription from the expression vector pSG5hPPARγ (Riboprobe system, Promega) and quantified by competitive PCR over a wide range of concentrations (0.25–25 attomole (amol) added in the RT reaction). Standard curves obtained when assaying PPARγ total mRNA or PPARγ2 mRNA are shown in Fig. 2 C. The linearity (r = 0.99) and the slopes of the standard curves (0.98 and 1.11) indicated that the RT-competitive PCR is quantitative and that all the mRNA molecules are copied into cDNA during the RT step. The lower limit of the assay was about 0.05 amol of mRNA in the RT reaction, and the interassay variation of the RT-competitive PCR was 7% with six separated determinations of the same amount of PPARγ mRNA.
      Figure thumbnail gr2
      Figure 2RT-competitive PCR method to measure PPARγ mRNA levels. A, scheme highlighting the features of the vector pSG5hPPARγ2 and the vector derived from it, pBSCompPPARγ, which served to synthesize competitor DNA. The primer pairs used in the RT and PCR reactions as well as the different-sized amplicons obtained are indicated.B, typical analysis of the fluorescence-labeled PCR products on an automated fluorescence DNA sequencer using a denaturing 4% polyacrylamide gel electrophoresis. C, validation of the RT-competitive PCR assay and standard curves obtained when assaying PPARγ total mRNA or PPARγ2 mRNA. The linearity (r = 0.99) and the slopes of the standard curves (0.98 and 1.11) indicated that the RT-competitive PCR is really quantitative and that all the mRNA molecules are copied into cDNA during the RT step.

      Western Blot Analysis of PPARγ

      Cells and tissues were homogenized in a lysis buffer of PBS containing 1% Triton X-100 (Sigma). Tissues were homogenized in extraction buffer containing PBS and 1% Nonidet P-40 (Sigma), 0.5% sodium deoxycholate (Sigma), 0.1% SDS (Sigma). Fresh mixture protease inhibitor (ICN) was added (100 mg/ml AEBSF, 5 mg/ml EDTA, 1 mg/ml leupeptin, 1 mg/ml pepstatin). Protein extracts were obtained by centrifugation of the lysate at 4 °C, and concentration was measured with the Bio-Rad DC Protein colorimetric assay system.
      Protein (100 μg) was separated by SDS-PAGE, transferred to nitrocellulose membrane (Amersham Life Science, Inc.), and blocked overnight in blocking buffer (20 mm Tris, 100 mm NaCl, 1% Tween-20, 10% skim milk). Filters were first incubated for 4 h at room temperature with rabbit IgG anti-mPPARγ (10 mg/ml), raised against an N-terminal PPARγ peptide (amino acids 20–104), and next developed for 1 h at room temperature with a goat anti-rabbit IgG (whole molecule) peroxidase conjugate (Sigma) diluted at 1/500. The complex was visualized with 4-chloro-1-naphtol as reagent.

      Analysis of Promoter Activity

      To test the activity of the human PPARγ promoters several reporter constructs were made. A 1-kb fragment of PAC clone 8856 was isolated by PCR using the oligonucleotides LF-35 (binding antisense in the PPARγ2 5′-UTR) and the oligonucleotide LF-58 (binding sense at position -1000 of the PPARγ2), was sequenced, and was inserted intoEcoRV site of pBluescript (Stratagene, La Jolla, CA). After digestion of plasmid pBSγ2p1000 with SmaI andKpnI, the insert was cloned into the reporter vector pGL3 (Promega), creating the expression vector pGL3γ2p1000. To isolate the PPARγ1 promoter, an 8-kb EcoRI fragment, which hybridized with the oligonucleotide LF-2 (corresponding to the 5′-UTR of γ1), was cloned into pBluescript. Partial mapping and sequencing of this clone revealed the presence of a 3-kb fragment upstream of the transcription initiation site. To test for promoter activity, aSacI/XhoI digestion of this clone containing the 3-kb promoter was inserted in the same sites of pGL3, resulting in the final vector pGL3γ1p3000. The pSG5-haPPARγ (
      • Aperlo C.
      • Pognonec P.
      • Saladin R.
      • Auwerx J.
      • Boulukos K.
      ) and pMSV-C/EBPα (
      • Christy R.J.
      • Yang V.W.
      • Ntambi J.M.
      • Geiman D.E.
      • Landschulz W.H.
      • Friedman A.D.
      • Nakabeppu Y.
      • Kelly T.J.
      • Lane M.D.
      ) expression vectors were described elsewhere. Transfections were carried out in 60-mm plates using standard calcium phosphate precipitation techniques (for 3T3-L1, CV-1, and COS cells) (
      • Schoonjans K.
      • Watanabe M.
      • Suzuki H.
      • Mahfoudi A.
      • Krey G.
      • Wahli W.
      • Grimaldi P.
      • Staels B.
      • Yamamoto T.
      • Auwerx J.
      ). Luciferase and β-galactosidase assays were carried out exactly as described previously (
      • Schoonjans K.
      • Watanabe M.
      • Suzuki H.
      • Mahfoudi A.
      • Krey G.
      • Wahli W.
      • Grimaldi P.
      • Staels B.
      • Yamamoto T.
      • Auwerx J.
      ).

      Electrophoretic Mobility Shift Assays (EMSA) and Oligonucleotide Sequences

      haPPARγ (
      • Aperlo C.
      • Pognonec P.
      • Saladin R.
      • Auwerx J.
      • Boulukos K.
      ), hPPARγ2, and mRXRα (
      • Leid M.
      • Kastner P.
      • Lyons R.
      • Nakshatri H.
      • Saunders M.
      • Zacharewski T.
      • Chen J.Y.
      • Staub A.
      • Garnier J.M.
      • Mader S.
      • Chambon P.
      ) proteins were synthesized in vitro in rabbit reticulocyte lysate (Promega). Molecular weights and quality of the in vitrotranslated proteins were verified by SDS-PAGE. PPAR (2 μl) and/or RXR (2 μl) were incubated for 15 min on ice in a total volume of 20 μl with 1-ng probe, 2.5 μg of poly(dI-dC) and 1 μg of herring sperm DNA in binding buffer (10 mm Tris-HCl pH 7.9, 40 mm KCl, 10% glycerol, 0.05% Nonidet P-40, and 1 mm dithiothreitol). For competition experiments, increasing amounts (from 10- to 200-fold molar excess) of cold oligonucleotide (AII-J-PPRE, 5′-GATCCTTCAACCTTTACCCTGGTAGA-3′ (
      • Vu-Dac N.
      • Schoonjans K.
      • Kosykh V.
      • Dallongeville J.
      • Fruchart J.-C.
      • Staels B.
      • Auwerx J.
      ); acyl-CoA oxidase (ACO)-PPRE, 5′-GATCCCGAACGTGACCTTTGTCCTGGTCCC-3′ (
      • Tugwood J.D.
      • Isseman I.
      • Anderson R.G.
      • Bundell K.R.
      • McPheat W.L.
      • Green S.
      ); or LPL-PPRE, 5′-GATCCGTCTGCCCTTTCCCCCTCTTCA-3′) (
      • Schoonjans K.
      • Peinado-Onsurbe J.
      • Heyman R.
      • Briggs M.
      • Cayet D.
      • Deeb S.
      • Staels B.
      • Auwerx J.
      ) were included just before adding T4-PNK end-labeled AII-J-PPRE oligonucleotide. DNA-protein complexes were separated by electrophoresis on a 4% polyacrylamide gel in 0.25 × TBE buffer at 4 °C (
      • Fried M.G.
      • Crothers D.M.
      ).

      DISCUSSION

      Two important findings recently underlined the importance of the PPARγ transcription factor. First, PPARγ has been identified as one of the key factors controlling adipocyte differentiation and function in rodent systems (
      • Tontonoz P.
      • Hu E.
      • Graves R.A.
      • Budavari A.I.
      • Spiegelman B.M.
      ,
      • Tontonoz P.
      • Hu E.
      • Spiegelman B.M.
      ). Second, the recent identification of prostaglandin J2 derivatives and antidiabetic thiazolidinediones as natural and synthetic PPARγ ligands, respectively (
      • Kliewer S.A.
      • Lenhard J.M.
      • Willson T.M.
      • Patel I.
      • Morris D.C.
      • Lehman J.M.
      ,
      • Forman B.M.
      • Tontonoz P.
      • Chen J.
      • Brun R.P.
      • Spiegelman B.M.
      • Evans R.M.
      ,
      • Lehmann J.M.
      • Moore L.B.
      • Smith-Oliver T.A.
      • Wilkison W.O.
      • Willson T.M.
      • Kliewer S.A.
      ,
      • Berger J.
      • Bailey P.
      • Biswas C.
      • Cullinan C.A.
      • Doebber T.W.
      • Hayes N.S.
      • Saperstein R.
      • Smith R.G.
      • Leibowitz M.D.
      ,
      • Willson T.M.
      • Cobb J.E.
      • Cowan D.J.
      • Wiethe R.W.
      • Correa I.D.
      • Prakash S.R.
      • Beck K.D.
      • Moore L.B.
      • Kliewer S.A.
      • Lehmann J.M.
      ). Thiazolidinediones are a new group of anti-diabetic drugs which improve insulin-resistance (for review, see Refs.
      • Hulin B.
      • McCarthy P.A.
      • Gibbs E.M.
      and
      • Saltiel A.R.
      • Olefsky J.M.
      ). The identification of thiazolidinediones as PPARγ ligands together with the central role that adipose tissue plays in the pathogenesis of important metabolic disorders, such as obesity and non-insulin-dependent diabetes mellitus (NIDDM), have generated a major interest to determine the role of this PPAR subtype in normal and abnormal adipocyte function in humans.
      The PPARγ gene spans about 100 kb and is composed of 9 exons, which give rise to PPARγ1 and PPARγ2 mRNAs by differential promoter usage and differential splicing. The gene structure as well as the sequence of the encoded protein are well conserved between human and mice (
      • Zhu Y.
      • Qi C.
      • Korenberg J.R.
      • Chen X.-N.
      • Noya D.
      • Rao M.S.
      • Reddy J.K.
      ) (99% similarity and 95% identity). Relative to the mouse, hamster, and Xenopus PPARγ (
      • Tontonoz P.
      • Hu E.
      • Graves R.A.
      • Budavari A.I.
      • Spiegelman B.M.
      ,
      • Aperlo C.
      • Pognonec P.
      • Saladin R.
      • Auwerx J.
      • Boulukos K.
      ,
      • Dreyer C.
      • Krey G.
      • Keller H.
      • Givel F.
      • Helftenbein G.
      • Wahli W.
      ), the human protein contains two additional amino acids. This is in agreement with the previous reports on the human PPARγ cDNA (
      • Elbrecht A.
      • Chen Y.
      • Cullinan C.A.
      • Hayes N.
      • Leibowitz M.D.
      • Moller D.E.
      • Berger J.
      ,
      • Lambe K.G.
      • Tugwood J.D.
      ,
      • Greene M.E.
      • Blumberg B.
      • McBride O.W.
      • Yi H.F.
      • Kronquist K.
      • Kwan K.
      • Hsieh L.
      • Greene G.
      • Nimer S.D.
      ). The availability of the structure of the human PPARγ gene and protein will now allow for genetic studies, evaluating its role in disorders such as insulin resistance, NIDDM, and diseases characterized by altered adipose tissue function such as obesity or lipodystrophic syndromes.
      To determine tissue-specific patterns of expression of the human PPARγ gene, we developed an RT-competitive PCR assay. Unlike results of previous reports, which used commercially available blots or single RNA samples (
      • Elbrecht A.
      • Chen Y.
      • Cullinan C.A.
      • Hayes N.
      • Leibowitz M.D.
      • Moller D.E.
      • Berger J.
      ,
      • Lambe K.G.
      • Tugwood J.D.
      ), we used multiple independent samples to base our conclusions on. As was observed in rodents (
      • Tontonoz P.
      • Hu E.
      • Graves R.A.
      • Budavari A.I.
      • Spiegelman B.M.
      ,
      • Tontonoz P.
      • Hu E.
      • Spiegelman B.M.
      ), we found PPARγ to be strongly expressed in adipose tissue. In addition to adipose tissue, the large intestine had high levels of PPARγ expression. Several other tissues, such as liver, kidney, and small intestine contained lower but nevertheless considerable levels of PPARγ RNA. Skeletal muscle, in contrast, contained only trace amounts of PPARγ mRNA.
      In adipose tissue and liver, about 15% of all PPARγ mRNA was of the PPARγ2 type, whereas in the remaining tissues no PPARγ2 mRNA was detected. These observations have several important implications. First, our data question the relative importance of PPARγ2. Indeed, our results in humans as well as the data by Xue et al. (
      • Xue J.C.
      • Schwarz E.J.
      • Chawla A.
      • Lazar M.A.
      ) in rodent adipocytes show consistently lower levels of PPARγ2 mRNA and protein relative to the PPARγ1 subtype. These observations are in line with the previous observations that the N-terminal domain of PPARγ was dispensable, both regarding transcriptional activation and capacity to induce adipocyte differentiation in vitro (
      • Tontonoz P.
      • Hu E.
      • Spiegelman B.M.
      ). However, the N-terminal domain is highly conserved between different species, suggesting it might have an important function in vivo. Second, PPARγ expression is much more widespread than previously realized, which implies that PPARγ controls gene expression in several tissues in addition to adipose tissue. Especially striking is the high level of PPARγ expression in the human large intestine. These reports are consistent with the reported high level expression of PPARγ in colonic mucosa in mouse (
      • Mansen A.
      • Guardiola-Diaz H.
      • Rafter J.
      • Branting C.
      • Gustafsson J.A.
      ). It is interesting to note that fatty acids, potential PPAR activators, have been shown to play an important role in modulating the function of the large intestine. For instance diets enriched in saturated lipids have been shown to predispose to the development of colon cancer (
      • Giovanucci E.
      • Willet W.C.
      ). Furthermore, it has been shown that diets enriched in ω-3 fatty acids, powerful PPAR activators, have a beneficial response on inflammatory diseases of the gastrointestinal tract such as colitis ulcerosa and Crohn's disease (
      • Stenson W.F.
      • Cort D.
      • Rodgers J.
      • Burakoff R.
      • DeSchryver-Kecskemeti K.
      • Gramlich T.L.
      • Beeken W.
      ,
      • Belluzi A.
      • Brignola C.
      • Campieri M.
      • Pera A.
      • Boschi S.
      • Miglioli M.
      ). Since the high level expression of PPARγ suggest that it might play an important role in normal and abnormal colonic function, further studies aimed at exploring this are definitely needed.Finally, the low levels of PPARγ expression in skeletal muscle cells also deserve some reflection. Muscle is responsible for clearance of the majority of glucose in the body and abnormal muscle glucose uptake is one of the prime features of insulin resistance and NIDDM. The low levels of PPARγ in muscle argue, therefore, that the beneficial effects of thiazolidinedione antidiabetic agents are not likely to be due to a direct effect of these agents on PPARγ present in the muscle. In fact, even though the liver has considerably higher levels of PPARγ relative to muscle, thiazolidinediones do not seem to affect PPAR responsive genes in liver tissue at the concentrations commonly used to lower glucose levels (
      • Schoonjans K.
      • Peinado-Onsurbe J.
      • Heyman R.
      • Briggs M.
      • Cayet D.
      • Deeb S.
      • Staels B.
      • Auwerx J.
      ). This observation together with the observed tissue distribution of PPARγ suggests that the glucose lowering effects of the thiazolidinedione PPARγ ligands are primarily a result of their activity on adipose tissue, which then, via a secreted signal, might influence muscle glucose uptake.
      To identify the molecular circuitry underlying tissue-specific expression of PPAR, we cloned and performed an initial characterization of the human PPARγ promoters. As shown, 3000 bp of the PPARγ1 and 1000 bp of the PPARγ2 promoter account for substantial levels of basal promoter activity. Further functional studies are underway to determine elements necessary for tissue-specific and regulated expression of the PPARγ gene. In this context, it will be interesting to determine the effects of transcription factors known to induce adipocyte differentiation on PPARγ expression in this tissue and to define the hierachical role that PPARγ plays in this process. PPARγ is not the only transcription factor involved in adipocyte differentiation. In addition to PPARγ, the basic helix-loop-helix leucine zipper factor ADD-1/SREBP1 and transcription factors of the C/EBP family also play a role in determining adipocyte differentiation. It is interesting to note that, as in the mouse PPARγ2 promoter (
      • Zhu Y.
      • Qi C.
      • Korenberg J.R.
      • Chen X.-N.
      • Noya D.
      • Rao M.S.
      • Reddy J.K.
      ), a potential consensus C/EBP response element could be identified in the human PPARγ2 promoter by homology searches. This observation fits well with the previous observation that forced expression of C/EBPβ could induce PPARγ expression and further studies on this subject are underway (
      • Wu Z.
      • Xie Y.
      • Bucher N.L.R.
      • Farmer S.R.
      ,
      • Wu Z.
      • Bucher N.L.R.
      • Farmer S.R.
      ).
      In conclusion, we report the characterization of the human PPARγ gene structure and furthermore define the structure of the PPARγ1 and γ2 promoter. In addition, our data show that human PPARγ has a similar structure and similar transactivation function as the rodent PPARs. The expression patterns of PPARγ1 and γ2 show that in man, PPARγ1 is the predominant form. Our results furthermore demonstrate that, in addition to adipose tissue, human colon expresses high levels of PPARγ. It is expected that the gene structure will facilitate our analysis of eventual PPARγ mutations in humans, whereas knowledge of expression patterns and sequence elements, as well as factors regulating PPARγ gene expression, could be of major importance in understanding PPAR biology.

      Acknowledgments

      The technical help of Delphine Cayet, Veerle Beelen, and Odile Vidal and the support and/or discussion with Drs. Rich Heyman and James Paterniti from Ligand Pharmaceuticals are kindly acknowledged. We acknowledge the gift of materials from Drs. Ronald Evans and Alex Nadzan. Access to an automatic sequencer by Drs. Philippe Froguel and Jorg Hager in the initial phases of this project are greatly appreciated.

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