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• W2093079433 abstract "In liver, the glyoxylate cycle contributes to two metabolic functions, urea and glucose synthesis. One of the key enzymes in this pathway is glyoxylate reductase/hydroxypyruvate reductase (GRHPR) whose dysfunction in human causes primary hyperoxaluria type 2, a disease resulting in oxalate accumulation and formation of kidney stones. In this study, we provide evidence for a transcriptional regulation by the peroxisome proliferator-activated receptor α (PPARα) of the mouse GRHPR gene in liver. Mice fed with a PPARα ligand or in which PPARα activity is enhanced by fasting increase their GRHPR gene expression via a peroxisome proliferator response element located in the promoter region of the gene. Consistent with these observations, mice deficient in PPARα present higher plasma levels of oxalate in comparison with their wild type counterparts. As expected, the administration of a PPARα ligand (Wy-14,643) reduces the plasma oxalate levels. Surprisingly, this effect is also observed in null mice, suggesting a PPARα-independent action of the compound. Despite a high degree of similarity between the transcribed region of the human and mouse GRHPR gene, the human promoter has been dramatically reorganized, which has resulted in a loss of PPARα regulation. Overall, these data indicate a species-specific regulation by PPARα of GRHPR, a key gene of the glyoxylate cycle. In liver, the glyoxylate cycle contributes to two metabolic functions, urea and glucose synthesis. One of the key enzymes in this pathway is glyoxylate reductase/hydroxypyruvate reductase (GRHPR) whose dysfunction in human causes primary hyperoxaluria type 2, a disease resulting in oxalate accumulation and formation of kidney stones. In this study, we provide evidence for a transcriptional regulation by the peroxisome proliferator-activated receptor α (PPARα) of the mouse GRHPR gene in liver. Mice fed with a PPARα ligand or in which PPARα activity is enhanced by fasting increase their GRHPR gene expression via a peroxisome proliferator response element located in the promoter region of the gene. Consistent with these observations, mice deficient in PPARα present higher plasma levels of oxalate in comparison with their wild type counterparts. As expected, the administration of a PPARα ligand (Wy-14,643) reduces the plasma oxalate levels. Surprisingly, this effect is also observed in null mice, suggesting a PPARα-independent action of the compound. Despite a high degree of similarity between the transcribed region of the human and mouse GRHPR gene, the human promoter has been dramatically reorganized, which has resulted in a loss of PPARα regulation. Overall, these data indicate a species-specific regulation by PPARα of GRHPR, a key gene of the glyoxylate cycle. As a major survival strategy, animals have developed metabolic pathways to store energy when food is abundant, which allows them to overcome periods of deprivation. Thus, they are able to switch from efficiently storing excess energy to its rapid mobilization to keep the organism alive when food is not available. Hormonal changes during fasting result in enhanced triglyceride hydrolysis in the adipose tissue, as well as glucose and ketone body production in the liver, to respond to the energy needs of the peripheral organs. Peroxisome proliferator-activated receptors (PPARs), 1The abbreviations used are: PPAR, peroxisome proliferator-activated receptor; AGT, alanine:glyoxylate aminotransferase; GFP, green fluorescent protein; ACO, acyl-CoA oxidase; FIAF, fasting-induced adipose factor; Luc, luciferase; h, human; m, mouse; RACE, rapid amplification of cDNA ends; ChIP, chromatin immunoprecipitation; UTR, untranslated region; PPRE, peroxisome proliferator response element; GRHPR, glyoxylate reductase/hydroxypyruvate reductase; RPA, RNase protection assay; RT, reverse transcription. as members of the nuclear hormone receptor superfamily, are involved in the transcriptional control of these metabolic pathways, including β-oxidation, gluconeogenesis, and amino acid catabolism, which underscores their importance in energy homeostasis (1Desvergne B. Wahli W. Endocr. Rev. 1999; 20: 649-688Crossref PubMed Scopus (2746) Google Scholar, 2Kersten S. Mandard S. Escher P. Gonzalez F.J. Tafuri S. Desvergne B. Wahli W. FASEB J. 2001; 15: 1971-1978Crossref PubMed Scopus (181) Google Scholar, 3Shearer B.G. Hoekstra W.J. Curr. Med. Chem. 2003; 10: 267-280Crossref PubMed Scopus (101) Google Scholar, 4Patsouris D. Mandard S. Voshol P.J. Escher P. Tan N.S. Havekes L.M. Koenig W. Marz W. Tafuri S. Wahli W. Muller M. Kersten S. J. Clin. Investig. 2004; 114: 94-103Crossref PubMed Scopus (215) Google Scholar). Upon activation by fatty acids and derivatives, they heterodimerize with the receptor for 9-cis-retinoid acid (retinoid X receptor, NR2B) and bind to peroxisome proliferator response elements (PPREs) in the promoter region of the genes whose expression they regulate. Three PPAR isotypes have been identified: PPARα (NR1C1); PPARβ/δ (NR1C2); and PPARγ (NR1C3). They exhibit different expression patterns and functions in animals (1Desvergne B. Wahli W. Endocr. Rev. 1999; 20: 649-688Crossref PubMed Scopus (2746) Google Scholar). PPARγ plays an important role in inflammation, lipid storage, and glucose homeostasis, whereas PPARβ/δ is important for skin functions, brain, and placenta development and also for energy homeostasis (3Shearer B.G. Hoekstra W.J. Curr. Med. Chem. 2003; 10: 267-280Crossref PubMed Scopus (101) Google Scholar). PPARα, which is a part of this study, regulates peroxisomal and mitochondrial fatty acid oxidation, microsomal fatty acid hydroxylation, lipoprotein metabolism, bile and amino acid metabolism, glucose homeostasis, biotransformation, inflammation, hepatocarcinogenesis in rodents, and other pathways and processes (5Mandard S. Muller M. Kersten S. Cell Mol. Life Sci. 2004; 61: 393-416Crossref PubMed Scopus (813) Google Scholar). In particular, this PPAR has been implicated in the regulation of the expression of two enzymes involved in the glyoxylate pathway, namely alanine:glyoxylate aminotransferase (AGT) and glyoxylate reductase/hydroxypyruvate reductase (GRHPR) (2Kersten S. Mandard S. Escher P. Gonzalez F.J. Tafuri S. Desvergne B. Wahli W. FASEB J. 2001; 15: 1971-1978Crossref PubMed Scopus (181) Google Scholar). AGT has a dual function, as alanine:glyoxylate aminotransferase and as serine: pyruvate aminotransferase, and is responsible for the conversion of glyoxylate into glycine and for the conversion of serine to hydroxypyruvate, respectively (see Fig. 1). Similarly, GRHPR functions both as glyoxylate reductase and as hydroxypyruvate reductase. GRHPR plays a key role in directing the carbon flux to gluconeogenesis by its ability to convert hydroxypyruvate into d-glycerate (see Fig. 1) (6Holmes R.P. Assimos D.G. J. Urol. 1998; 160: 1617-1624Crossref PubMed Scopus (146) Google Scholar). Therefore, regulation of this enzyme by PPARα may contribute to the function of the receptor in energy homeostasis. Linked to their role in metabolism, AGT and GRHPR are associated to primary hyperoxaluria type 1 and type 2, respectively, which are caused by an overproduction of oxalate. Oxalate is an inorganic acid that, when combined with calcium, produces insoluble calcium oxalate, which is the most common constituent of kidney stones (7Ogawa Y. Miyazato T. Hatano T. World J. Surg. 2000; 24: 1154-1159Crossref PubMed Scopus (44) Google Scholar). Deficiency in these two enzymes results in glyoxylate accumulation (see Fig. 1) and consequently to oxalate overproduction (8Holmes R.P. Goodman H.O. Assimos D.G. Kidney Int. 2001; 59: 270-276Abstract Full Text Full Text PDF PubMed Scopus (371) Google Scholar, 9Leumann E. Hoppe B. J. Am. Soc. Nephrol. 2001; 12: 1986-1993PubMed Google Scholar). Moreover, a decrease in glycolate oxidase activity and an increase in lactate dehydrogenase activity were observed after clofibrate treatment in rats, which results in an increase in oxalate concentration in urines (10Sharma V. Schwille P.O. Metabolism. 1997; 46: 135-139Abstract Full Text PDF PubMed Scopus (7) Google Scholar). Therefore, we investigated the involvement of PPARα in the regulation of glyoxylate metabolism and consequently in oxalate production. Mice were housed in a temperature-controlled room (23 °C) on a 10-h dark, 14-h light cycle. Pure-bred wild type or PPARα knock-out (11Lee S.S. Pineau T. Drago J. Lee E.J. Owens J.W. Kroetz D.L. Fernandez-Salguero P.M. Westphal H. Gonzalez F.J. Mol. Cell. Biol. 1995; 15: 3012-3022Crossref PubMed Scopus (1506) Google Scholar) mice of age 8-10 weeks were used for all of the experiments. Animal experiments were approved by the Service Veterinaire of the Canton de Vaud. Fasted animals were deprived of food for 24 h starting at the beginning of the light cycle. The PPARα ligand Wy-14,643 (50 mg/kg/day) or vehicle (0.5% carboxymethyl-cellulose) were administrated by gavage for 5 days (RNase protection assay (RPA) chromatin immunoprecipitation (ChIP)) or mixed to the food (0.1%) for 5 days (oxalate assay). Full-length mouse GRHPR cDNA was amplified using the following primer: 5′-ACCGCTCGAGCATGAAACCGGCGCGAC-3′ and 5′-GGGGTACCTTACAGCTTGAGTTC-3′. The amplification product was digested with XhoI/KpnI and subcloned into pEGFP-N2 or pEGFP-C2 (BD Biosciences). The mouse promoter region was amplified from a λ-phage containing a genomic fragment of the mouse promoter region. The different fragments of the promoter were obtained using the common downstream primer (5′-GGGGTACCCCAAGACCCGGAGCAGCAAACA-3′) and three different upstream primers (5′-TCCCCCGGGGGAGGTTGTAAGCCACCATTTGGT-3′, 5′-TCCCCCGGGGGACTGGTGGGGATATGTGTTT-3′, and 5′-TCCCCCGGGGGATCTACCCGTGGCTCAGCATA-3′) for amplification and subcloned in plasmids p2500-Luc, p1100-Luc, and p400-Luc, respectively. Similarly, the human promoter was amplified from genomic DNA extracted from HepG2 cells. A common downstream primer (5′-CCGCTCGAGCGGCATGAGTCGCACCGGTCTCATC-3′) and the upstream primers (5′-GGGGTACCCCTCAAGGAAACCAACCCTGGTGC-3′, 5′-GGGGTCCCCCGGGACTCAGCCACCAC-AACCA-3′, and 5′-GGGGTACCCCGAGGCGGGAGGATCAT-TGGAGCAC-3′) were used for the amplification of the fragments subcloned in plasmids p4500-Luc, p3000-Luc, and p2000-Luc, respectively. PCR was performed using the following conditions: 95 °C for 4 min; 35 cycles at 94 °C for 45 s; 55 °C for 45 s; and 72 °C for 2 min with a final elongation step at 72 °C for 7 min. PCR products were digested with KpnI and SmaI for the mouse fragments and with KpnI and XhoI for the human fragments and subcloned in front of the luciferase reporter gene into the ΔpGL2 basic vector (Promega). HepG2 cells were cultured in minimum essential medium (Sigma) supplemented with 10% fetal calf serum (Hyclone), 1 × minimal essential medium non-essential amino acids (Sigma), 1 mm sodium pyruvate (Sigma), and 1% penicillin/streptomycin (Invitrogen). 1 × 105 cells/well were seeded in 12-well tissue culture plates the day before transfection. For cotransfections, 1.5 μg of reporter vector, 0.1 μg of PPARα (pCDNA3.1/hPPARα), and 0.4 μg of pEF1/Myc-His expressing β-galactosidase (Invitrogen) were used for each well. The total amount of DNA was adjusted with pCDNA3.1 to 3 μg. Transfections were performed using the Superfect™ Transfection Reagent (Qiagen) according to the manufacturer's instructions. 5 h after transfection, serum was removed and cells were treated with 10 μm Wy-14,643 (ChemSyn Laboratories) or Me2SO. After 24 h, cells were washed with phosphate buffer and lysed in reporter lysis buffer (Promega). For the β-galactosidase assay, lysis solution was mixed with 2× assay buffer (200 mm sodium phosphate buffer, pH 7.3, 2 mm MgCl2; 100 mm β-mercaptoethanol, 1.33 mg/ml 2-nitrophenyl β-d-galactopyranoside). The solution was incubated for 5 min at 37 °C, and the absorbance was measured at 420 nm. Luciferase was measured using the luciferase assay system (Promega). γ-32P-End-labeled primers 5′-CATGAGTCGCGCCGGTTTCATAAGAC-3′ (mouse) and 5′-ACCTGGCAGTACAGAAGCTGGC-3′ (human) were used for reverse transcription (RT) and sequencing. The sequencing was carried out using the fmol DNA cycle sequencing system (Promega). 50 μg of total RNA was mixed with the labeled primer (3 μm) in hybridization buffer (150 mm KCl, 10 mm Tris, pH 8.3, 1 mm EDTA). The mixture was heated at 95 °C for 2 min and then placed at 65 °C for 1 h and 30 min at room temperature for 1 h and 30 min and finally at 4 °C overnight. Reverse transcription was then performed with the Superscript II RNase H-reverse transcriptase (Invitrogen). Total RNA was isolated using the TRIzol reagent (Invitrogen). Northern blot analysis was performed using 30 μg of total RNA according to standard protocols (12Sambrook J. Fritsch E.F. Maniatis T. Molecular Cloning: a Laboratory Manual. 2nd Ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY1989Google Scholar). Mouse GRHPR cDNA probe was random-primed-labeled using the High Prime kit (Roche Applied Science). RT-PCR/RPA—Gene-specific probes for the mouse GRHPR, AGT, and lactate dehydrogenase A were obtained by RT-PCR from mouse liver total RNA and cloned into the pGEM T-Easy vector (Promega). RT-PCR was performed with the Titan One Tube RT-PCR system (Roche Applied Science). 100 ng of total RNA was used for each PCR. The number of cycles was first determined to find the exponential phase of each set of primers. Gene-specific antisense riboprobes were synthesized by in vitro transcription with either T7 or Sp6 RNA polymerase (Ambion). For all of the riboprobes with the exception of L27, a ratio of 1:1 of [α-32P]UTP to cold UTP was used, whereas a ratio of 1:20 was used for L27 probe. RPA was carried out using the Direct Protect lysate RPA kit (Ambion) with the following modifications. 15 μg of total RNA were resuspended in 45 μl of lysis buffer and incubated with 5 × 104 cpm of gene-specific and L27 riboprobes. RPA products were resolved in a 6% electrolyte gradient-denaturing polyacrylamide gel. Gels were dried and exposed to x-ray film. Pure-bred wild type or PPARα null mice on a sv129 background were used. Mice were fed by gavage with either Wy-14,643 (PPARα ligand; 50 mg/kg/day) or vehicle (0.5% carboxymethylcellulose) for 5 days. After the indicated treatment, mice were sacrificed by cervical dislocation. The liver was rapidly perfused with prewarmed (37 °C) phosphate-buffered saline for 5 min followed by 0.2% collagenase for 10 min. The liver then was diced and forced through a 60 μm stainless steel sieve, and the hepatocytes were collected directly into Dulbecco's modified Eagle's medium containing 1% formaldehyde. After incubation at 37 °C for 15 min, the hepatocytes were pelleted and ChIP was performed using PPARα antibody as previously described (13Di Poi N. Tan N.S. Michalik L. Wahli W. Desvergne B. Mol. Cell. 2002; 10: 721-733Abstract Full Text Full Text PDF PubMed Scopus (290) Google Scholar). PCR was performed using primers flanking the mGRHPR PPREs. The primers flanking the mouse PPREs were as follows: 5′-CCCATGGGACAGATAAGGAAGACA-3′ and 5′-CCACCCAGGCGAGCTAGACACAA-G-3′ for PPRE1; 5′-AGGCTGGCCACAAACTCAC-TCT-3′ and 5′-CTGCCACCGGACCTTCATT-TT-3′ for PPRE2; and 5′-CTGGCCTGGGGACACGAAAAC-3′ and 5′-GGGCGCCAAGGACAA-CACAGT-3′ for the negative control. HepG2 cells were cultured in 10-cm dishes and transfected with a PPARα-expressing construct. 16 h post-transfection, cells were induced with either 0.01% Me2SO (vehicle) or 10 μm Wy-14,643 for 6 h and then fixed in 1% formaldehyde. After 15 min at 37 °C, cells were pelleted and ChIP was performed using a PPARα antibody as previously described (13Di Poi N. Tan N.S. Michalik L. Wahli W. Desvergne B. Mol. Cell. 2002; 10: 721-733Abstract Full Text Full Text PDF PubMed Scopus (290) Google Scholar). The PCR primers for the putative human PPREs were 5′-TTGCCAAGGACCCACTTTGTACTGAG-3′ and 5′-TGGACTGGGCCAGGAAAGATAAGGT-3′ for hPPRE3; and 5′-CCCTGTGAATGTGGGAAAGCTCTT-3′ and 5′-GGGAGGCCCTCAGGAGAAGCAGGA-3′ for hPPRE4; 5′-TCAAAGTCATCAGCACCATGTCTGTG-3′ and 5′-ATAACACCTGCCTTTGCTACTTCAAG-3′ for hPPRE5; and 5′-AAGTTCAGAGCTGGGAAGGCGAACAG-3′ and 5′-TAGAGGGAGAGGAGGCAGGGTTGAG-3′ for the fasting-induced adipose factor (FIAF)-positive control PPRE. Mouse plasma was acidified by dilution 1:1 with 0.15 n HCl to dissociate oxalate from plasmatic proteins. Plasmatic proteins were then removed by ultrafiltration and by centrifugation at 14,000 × g (1 h, room temperature) in a Microcon YM30 column (Amicon). The measure of oxalate in ultrafiltrate was done using the oxalate oxidase/peroxidase method (Kit Oxalate, Dialine) on a Cobas FARA automate (Roche Applied Science). Using SABRE (selective amplification via biotin and restriction-mediated enrichment) (14Lavery D.J. Lopez-Molina L. Fleury-Olela F. Schibler U. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 6831-6836Crossref PubMed Scopus (47) Google Scholar), we previously isolated the cDNA of a novel target gene for PPARα whose identity and regulation are studied herein. The full-length cDNA was amplified using 5′- and 3′-RACE PCR, cloned, and sequenced. A comparison of these mouse cDNA and derived protein sequences with those present in the NCBI data base indicated a very high similarity with the human GRHPR sequence. This analysis also revealed the presence of a region presenting a high homology with the d-isomer 2-hydroxyacid dehydrogenase consensus motif (PROSITE accession number PS00671) and a putative binding site for nicotinamide adenine dinucleotide (NAD). These characteristics suggested that the gene encodes a protein with an enzymatic activity, most likely mouse GRHPR. The amino acid sequence analysis revealed a Leu-Lys-Leu motif at the protein C terminus, which is close to the consensus motif of the peroxisomal targeting signal-1 that typically consists of these three amino acids or a conservative variant thereof (15Gould S.J. Collins C.S. Nat. Rev. Mol. Cell. Biol. 2002; 3: 382-389Crossref PubMed Scopus (93) Google Scholar). This observation suggested that the cloned cDNA might encode a peroxisomal protein. To determine the subcellular localization of this gene product, fusion proteins with the green fluorescent protein (GFP) localized either at the C-terminal (GRHPR N2) or at the N-terminal (GRHPR C2) end of GRHPR were generated. When transfected into HEK 293 cells, the constructs produced cytoplasmic fusion proteins (Fig. 2). As a control, a GFP fusion acyl-CoA oxidase (ACO) was targeted to peroxisomes as expected, whereas the mutation of the peptide signal Ser-Lys-Leu into Leu-Lys-Leu, as found in GRHPR, resulted in a cytoplasmic localization of the mutated ACO. These results suggested that the cloned cDNA encodes a protein localized in the cytoplasm. We then examined the expression profile of the gene in different organs by Northern blot analysis. As shown in Fig. 3, high expression levels were detected in the liver and a much lower expression was also found in the kidney. Thus, based on the high percentage of identity, cytoplasmic localization, identical gene organization (see below), and expression profile in liver and kidney as human GRHPR (16Giafi C.F. Rumsby G. Ann. Clin. Biochem. 1998; 35: 104-109Crossref PubMed Scopus (68) Google Scholar), we concluded that the identified gene encodes the mouse GRHPR.Fig. 3Expression pattern of mouse GRHPR. Northern blot analysis of mouse total RNA (30 μg/lane) from various tissues was performed using a GRHPR-specific probe. The mRNA level of the L27 ribosomal protein (L27) was used as control.View Large Image Figure ViewerDownload Hi-res image Download (PPT) We then investigated the genomic organization of this GRHPR gene. Genomic DNA was obtained by screening a mouse λ-phage genomic library using two probes: one from the 5′ end (nucleotides 40-370) and one from the 3′ end (nucleotides 938-1237) of the cDNA. Two overlapping clones were identified containing the entire gene and ~10 kb of DNA upstream of the translation initiation codon ATG (data not shown). After sequencing, a comparison of the cDNA and the genomic sequences allowed determination of the organization of the gene (Fig. 4A), which was recently mapped on mouse chromosome 4 B2 (17Waterston R.H. Lindblad-Toh K. Birney E. Rogers J. Abril J.F. Agarwal P. Agarwala R. Ainscough R. Alexandersson M. An P. Antonarakis S.E. Attwood J. Baertsch R. Bailey J. Barlow K. Beck S. Berry E. Birren B. Bloom T. Bork P. Botcherby M. Bray N. Brent M.R. Brown D.G. Brown S.D. Bult C. Burton J. Butler J. Campbell R.D. Carninci P. Cawley S. Chiaromonte F. Chinwalla A.T. Church D.M. Clamp M. Clee C. Collins F.S. Cook L.L. Copley R.R. Coulson A. Couronne O. Cuff J. Curwen V. Cutts T. Daly M. David R. Davies J. Delehaunty K.D. Deri J. Dermitzakis E.T. Dewey C. Dickens N.J. Diekhans M. Dodge S. Dubchak I. Dunn D.M. Eddy S.R. Elnitski L. Emes R.D. Eswara P. Eyras E. Felsenfeld A. Fewell G.A. Flicek P. Foley K. Frankel W.N. Fulton L.A. Fulton R.S. Furey T.S. Gage D. Gibbs R.A. Glusman G. Gnerre S. Goldman N. Goodstadt L. Grafham D. Graves T.A. Green E.D. Gregory S. Guigo R. Guyer M. Hardison R.C. Haussler D. Hayashizaki Y. Hillier L.W. Hinrichs A. Hlavina W. Holzer T. Hsu F. Hua A. Hubbard T. Hunt A. Jackson I. Jaffe D.B. Johnson L.S. Jones M. Jones T.A. Joy A. Kamal M. Karlsson E.K. Karolchik D. Kasprzyk A. Kawai J. Keibler E. Kells C. Kent W.J. Kirby A. Kolbe D.L. Korf I. Kucherlapati R.S. Kulbokas E.J. Kulp D. Landers T. Leger J.P. Leonard S. Letunic I. Levine R. Li J. Li M. Lloyd C. Lucas S. Ma B. Maglott D.R. Mardis E.R. Matthews L. Mauceli E. Mayer J.H. McCarthy M. McCombie W.R. McLaren S. McLay K. McPherson J.D. Meldrim J. Meredith B. Mesirov J.P. Miller W. Miner T.L. Mongin E. Montgomery K.T. Morgan M. Mott R. Mullikin J.C. Muzny D.M. Nash W.E. Nelson J.O. Nhan M.N. Nicol R. Ning Z. Nusbaum C. O'Connor M.J. Okazaki Y. Oliver K. Overton-Larty E. Pachter L. Parra G. Pepin K.H. Peterson J. Pevzner P. Plumb R. Pohl C.S. Poliakov A. Ponce T.C. Ponting C.P. Potter S. Quail M. Reymond A. Roe B.A. Roskin K.M. Rubin E.M. Rust A.G. Santos R. Sapojnikov V. Schultz B. Schultz J. Schwartz M.S. Schwartz S. Scott C. Seaman S. Searle S. Sharpe T. Sheridan A. Shownkeen R. Sims S. Singer J.B. Slater G. Smit A. Smith D.R. Spencer B. Stabenau A. Stange-Thomann N. Sugnet C. Suyama M. Tesler G. Thompson J. Torrents D. Trevaskis E. Tromp J. Ucla C. Ureta-Vidal A. Vinson J.P. Von Niederhausern A.C. Wade C.M. Wall M. Weber R.J. Weiss R.B. Wendl M.C. West A.P. Wetterstrand K. Wheeler R. Whelan S. Wierzbowski J. Willey D. Williams S. Wilson R.K. Winter E. Worley K.C. Wyman D. Yang S. Yang S.P. Zdobnov E.M. Zody M.C. Lander E.S. Nature. 2002; 420: 520-562Crossref PubMed Scopus (5442) Google Scholar). The gene spans over ~10 kb and is composed of 9 exons. The different exon lengths range from 73 (Exon 3) to 321 bp (Exon 9) in size corresponding to a total coding region of 987 bp for a transcribed region of 9369 bp (see below). The ATG start codon was located in Exon 1, 80 bp downstream of the transcription initiation site (see below). The last exon, Exon 9, contains the stop codon TAA and a 3′-UTR (3′-untranslated region) of 199 bp in which a polyadenylation signal, AUUAAA, was found 19 bp upstream of the poly(A) tail. The eight introns are ranging in size from 489 bp (Intron III) to 1568 bp (Intron VII). All of the exon-intron boundaries have the canonical GT/AG motif with the exception of the splice donor of Exon 2, which has a non-canonical site TA/AG (Fig. 4B). A comparison with the human GRHPR gene localized on chromosome 9q12 (18Huang T. Yang W. Pereira A.C. Craigen W.J. Shih V.E. Biochem. Biophys. Res. Commun. 2000; 268: 298-301Crossref PubMed Scopus (6) Google Scholar) revealed an identical genomic organization with eight introns flanked by nine exons. The location of the eight introns is strictly conserved, and the length of the coding sequence of the human gene is the same (987 bp) as that of the mouse gene (Fig. 4B). However, the length of the introns varies between the two species. Furthermore, the 5′-UTR (106 bp) and the 3′-UTR (207 bp) of the human transcript are longer than in the mouse (80 and 199 bp, respectively). Firstly, the transcription initiation site of the two genes was determined. Primer extension analysis showed that the transcription start sites were 24 bp upstream in mouse (Fig. 5A) and 26 bp upstream in human (Fig. 5B) of the 5′ends of cDNAs contained in GenBank™ data base (accession numbers BY353228 and CB998056). In human, two primer extension products separated by only 1 bp were detected, possibly reflecting 5′-cap methylation of the mRNA, which may cause incomplete reverse transcription of some of the mRNA molecules. Secondly, the alignment of the two promoter sequences showed a surprisingly poor homology at first sight but further analysis revealed that the proximal region of the mouse promoter (~-300 to ~-2000) is found in a reverse orientation in the human gene 4 kb upstream the transcription initiation site (~-3900 to ~-5600) (Fig. 5C). Interestingly, the functional PPRE in the mouse promoter (mPPRE1) (see below) was also found but less conserved in human in this shifted region (Fig. 5C). To know whether this displacement and inversion of promoter region was specific to human, the GRHPR promoter of other species was also analyzed. Comparison with the rat (Ensemble data base accession number RNOR03291619), dog (Ensemble data base contig 55728.1.64020), and chimpanzee (Ensemble data base accession number AADA01236509) promoter showed that the rearrangement is also present in the chimpanzee but not in the rat and dog promoter (Fig. 5D). The alignment in the same orientation of the rearranged promoter fragment from the mouse, rat, chimpanzee, and human promoter revealed conserved regions among the four species, which may suggest that regulatory functions have been maintained after reconfiguration of the promoter in primates (Fig. 6). Interestingly, however, the functional PPRE identified in the mouse sequence (see below) was not conserved in any of the other species, not even in the rat where the half-sites of the response element are separated by 12 nucleotides (Fig. 6). Further analysis of the promoter demonstrated a high level of sequence repeats in the human promoter between the transcription initiation site and the boundary of the inverted region. Indeed, this region contains 64% repeated elements (41.5% short interspeed elements, 16.5% long interspeed elements, and 6% long terminal repeat elements), whereas the average of the human genome is 46%. In comparison, only 21% of the mouse promoter region is constituted of such elements (21% short interspeed elements), which is under the average of 37.5% for the complete genome (17Waterston R.H. Lindblad-Toh K. Birney E. Rogers J. Abril J.F. Agarwal P. Agarwala R. Ainscough R. Alexandersson M. An P. Antonarakis S.E. Attwood J. Baertsch R. Bailey J. Barlow K. Beck S. Berry E. Birren B. Bloom T. Bork P. Botcherby M. Bray N. Brent M.R. Brown D.G. Brown S.D. Bult C. Burton J. Butler J. Campbell R.D. Carninci P. Cawley S. Chiaromonte F. Chinwalla A.T. Church D.M. Clamp M. Clee C. Collins F.S. Cook L.L. Copley R.R. Coulson A. Couronne O. Cuff J. Curwen V. Cutts T. Daly M. David R. Davies J. Delehaunty K.D. Deri J. Dermitzakis E.T. Dewey C. Dickens N.J. Diekhans M. Dodge S. Dubchak I. Dunn D.M. Eddy S.R. Elnitski L. Emes R.D. Eswara P. Eyras E. Felsenfeld A. Fewell G.A. Flicek P. Foley K. Frankel W.N. Fulton L.A. Fulton R.S. Furey T.S. Gage D. Gibbs R.A. Glusman G. Gnerre S. Goldman N. Goodstadt L. Grafham D. Graves T.A. Green E.D. Gregory S. Guigo R. Guyer M. Hardison R.C. Haussler D. Hayashizaki Y. Hillier L.W. Hinrichs A. Hlavina W. Holzer T. Hsu F. Hua A. Hubbard T. Hunt A. Jackson I. Jaffe D.B. Johnson L.S. Jones M. Jones T.A. Joy A. Kamal M. Karlsson E.K. Karolchik D. Kasprzyk A. Kawai J. Keibler E. Kells C. Kent W.J. Kirby A. Kolbe D.L. Korf I. Kucherlapati R.S. Kulbokas E.J. Kulp D. Landers T. Leger J.P. Leonard S. Letunic I. Levine R. Li J. Li M. Lloyd C. Lucas S. Ma B. Maglott D.R. Mardis E.R. Matthews L. Mauceli E. Mayer J.H. McCarthy M. McCombie W.R. McLaren S. McLay K. McPherson J.D. Meldrim J. Meredith B. Mesirov J.P. Miller W. Miner T.L. Mongin E. Montgomery K.T. Morgan M. Mott R. Mullikin J.C. Muzny D.M. Nash W.E. Nelson J.O. Nhan M.N. Nicol R. Ning Z. Nusbaum C. O'Connor M.J. Okazaki Y. Oliver K. Overton-Larty E. Pachter L. Parra G. Pepin K.H. Peterson J. Pevzner P. Plumb R. Pohl C.S. Poliakov A. Ponce T.C. Ponting C.P. Potter S. Quail M. Reymond A. Roe B.A. Roskin K.M. Rubin E.M. Rust A.G. Santos R. Sapojnikov V. Schultz B. Schultz J. Schwartz M.S. Schwartz S. Scott C. Seaman S. Searle S. Sharpe T. Sheridan A. Shownkeen R. Sims S. Singer J.B. Slater G. Smit A. Smith D.R. Spencer B. Stabenau A. Stange-Thomann N. Sugnet C. Suyama M. Tesler G. Thompson J. Torrents D. Trevaskis E. Tromp J. Ucla C. Ureta-Vidal A. Vinson J.P. Von Niederhausern A.C. Wade C.M. Wall M. Weber R.J. Weiss R.B. Wendl M.C. West A.P. Wetterstrand K. Wheeler R. Whelan S. Wierzbowski J. Willey D. Williams S. Wilson R.K. Winter E. Worley K.C. Wyman D. Yang S. Yang S.P. Zdobnov E.M. Zody M.C. Lander E.S. Nature. 2002; 420: 520-562Crossref PubMed Scopus (5442) Google Scholar). Interestingly, the percentage of repeats in the shifted region is 25 and 20% for the human and the mouse, respectively. These results suggest that even though the shifted region may have some regulatory function, the promoter region in primates underwent dramatic rearrangements also attested by its high content in repeated elements and might have lost PPAR responsiveness (see below). However, an analysis of the human and mouse 5′ region over 4500 bp revealed several putative PPREs (NUBIScan algorithm) (19P" @default.
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• W2093079433 title "Promoter Rearrangements Cause Species-specific Hepatic Regulation of the Glyoxylate Reductase/Hydroxypyruvate Reductase Gene by the Peroxisome Proliferator-activated Receptor α" @default.
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