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• W2093961921 abstract "Gluconeogenesis contributes to insulin resistance in type 1 and type 2 diabetes, but its regulation and the underlying molecular mechanisms remain unclear. Recently, calcium-regulated heat-stable protein 1 (CARHSP1) was identified as a biomarker for diabetic complications. In this study, we investigated the role of CARHSP1 in hepatic gluconeogenesis. We assessed the regulation of hepatic CARHSP1 expression under conditions of fasting and refeeding. Adenovirus-mediated CARHSP1 overexpression and siRNA-mediated knockdown experiments were performed to characterize the role of CARHSP1 in the regulation of gluconeogenic gene expression. Here, we document for the first time that CARHSP1 is regulated by nutrient status in the liver and functions at the transcriptional level to negatively regulate gluconeogenic genes, including the glucose-6-phosphatase catalytic subunit (G6Pc) and phosphoenolpyruvate carboxykinase 1 (PEPCK1). In addition, we found that CARHSP1 can physically interact with peroxisome proliferator-activated receptor-α (PPARα) and inhibit its transcriptional activity. Both pharmacological and genetic ablations of PPARα attenuate the inhibitory effect of CARHSP1 on gluconeogenic gene expression in hepatocytes. Our data suggest that CARHSP1 inhibits hepatic gluconeogenic gene expression via repression of PPARα and that CARHSP1 may be a molecular target for the treatment of diabetes. Gluconeogenesis contributes to insulin resistance in type 1 and type 2 diabetes, but its regulation and the underlying molecular mechanisms remain unclear. Recently, calcium-regulated heat-stable protein 1 (CARHSP1) was identified as a biomarker for diabetic complications. In this study, we investigated the role of CARHSP1 in hepatic gluconeogenesis. We assessed the regulation of hepatic CARHSP1 expression under conditions of fasting and refeeding. Adenovirus-mediated CARHSP1 overexpression and siRNA-mediated knockdown experiments were performed to characterize the role of CARHSP1 in the regulation of gluconeogenic gene expression. Here, we document for the first time that CARHSP1 is regulated by nutrient status in the liver and functions at the transcriptional level to negatively regulate gluconeogenic genes, including the glucose-6-phosphatase catalytic subunit (G6Pc) and phosphoenolpyruvate carboxykinase 1 (PEPCK1). In addition, we found that CARHSP1 can physically interact with peroxisome proliferator-activated receptor-α (PPARα) and inhibit its transcriptional activity. Both pharmacological and genetic ablations of PPARα attenuate the inhibitory effect of CARHSP1 on gluconeogenic gene expression in hepatocytes. Our data suggest that CARHSP1 inhibits hepatic gluconeogenic gene expression via repression of PPARα and that CARHSP1 may be a molecular target for the treatment of diabetes. Gluconeogenesis maintains glucose homeostasis in humans, especially during prolonged fasting or starvation, but abnormal hepatic gluconeogenesis contributes to insulin resistance (1Boden G. Chen X. Stein T.P. Am. J. Physiol. Endocrinol. Metab. 2001; 280: E23-30Crossref PubMed Google Scholar, 2Perriello G. Pampanelli S. Del Sindaco P. Lalli C. Ciofetta M. Volpi E. Santeusanio F. Brunetti P. Bolli G.B. Diabetes. 1997; 46: 1010-1016Crossref PubMed Scopus (77) Google Scholar, 3Wajngot A. Chandramouli V. Schumann W.C. Ekberg K. Jones P.K. Efendic S. Landau B.R. Metabolism. 2001; 50: 47-52Abstract Full Text PDF PubMed Scopus (111) Google Scholar) with increased gluconeogenesis as a major contributor to fasting hyperglycemia in both type 1 and type 2 diabetes (4Biddinger S.B. Kahn C.R. Annu. Rev. Physiol. 2006; 68: 123-158Crossref PubMed Scopus (581) Google Scholar, 5Quinn P.G. Yeagley D. Curr. Drug Targets Immune Endocr. Metabol. Disord. 2005; 5: 423-437Crossref PubMed Scopus (96) Google Scholar). Gluconeogenesis is controlled by certain rate-limiting enzymes such as the glucose-6-phosphatase catalytic subunit (G6Pc) 3The abbreviations used are: G6Pcglucose-6-phosphatase, catalytic subunitPEPCKphosphoenolpyruvate carboxykinasePEPCK1phosphoenolpyruvate carboxykinase 1 (soluble)LucluciferaseIPimmunoprecipitationDEXdexamethasonePPARαperoxisome proliferator-activated receptor αPPREPPAR-response elementTK-RLthymidine kinase promoter-renilla luciferasem.o.i.multiplicity of infectionaaamino acid. and phosphoenolpyruvate carboxykinase (PEPCK), and these genes are regulated by critical metabolism-related hormones including insulin, glucagon, and glucocorticoids. Although the important role of gluconeogenesis is known, the regulation of gluconeogenesis and its underlying mechanisms still remain to be further investigated. glucose-6-phosphatase, catalytic subunit phosphoenolpyruvate carboxykinase phosphoenolpyruvate carboxykinase 1 (soluble) luciferase immunoprecipitation dexamethasone peroxisome proliferator-activated receptor α PPAR-response element thymidine kinase promoter-renilla luciferase multiplicity of infection amino acid. Calcium-regulated heat-stable protein (CARHSP1) is a ubiquitously expressed phosphoprotein that is comprised of 147 amino acids with nearly 14% proline (6Groblewski G.E. Yoshida M. Bragado M.J. Ernst S.A. Leykam J. Williams J.A. J. Biol. Chem. 1998; 273: 22738-22744Abstract Full Text Full Text PDF PubMed Scopus (50) Google Scholar). CARHSP1 is serine-phosphorylated by Akt, SGK1 (serum- and glucocorticoid-induced protein kinase 1) and RSK (p90 ribosomal S6 kinase) in response to growth factors such as EGF and insulin-like growth factor-1 (IGF-1) (7Auld G.C. Campbell D.G. Morrice N. Cohen P. Biochem. J. 2005; 389: 775-783Crossref PubMed Scopus (25) Google Scholar). On the other hand, CARHSP1 can be dephosphorylated by Ca2+/calmodulin-regulated protein phosphatase calcineurin (PP2B) (8Schäfer C. Steffen H. Krzykowski K.J. Göke B. Groblewski G.E. Am. J. Physiol. Gastrointest. Liver Physiol. 2003; 285: G726-734Crossref PubMed Scopus (16) Google Scholar). Coordinated phosphorylation and dephosphorylation of CARHSP1 contribute to the multiple phosphorylated isoforms of CARHSP1 in acinar cells (9Lee S. Wishart M.J. Williams J.A. Biochem. Biophys. Res. Commun. 2009; 385: 413-417Crossref PubMed Scopus (6) Google Scholar). Recently, CARHSP1 was discovered as a biomarker for diabetic retinopathy (10Freeman W.M. Bixler G.V. Brucklacher R.M. Lin C.M. Patel K.M. Vanguilder H.D. Lanoue K.F. Kimball S.R. Barber A.J. Antonetti D.A. Gardner T.W. Bronson S.K. Pharmacogenomics J. 2009; 10: 385-395Crossref PubMed Scopus (26) Google Scholar) and as a regulator of TNF-α mRNA stability in macrophages (11Pfeiffer J.R. McAvoy B.L. Fecteau R.E. Deleault K.M. Brooks S.A. Mol. Cell. Biol. 2011; 31: 277-286Crossref PubMed Scopus (38) Google Scholar). CARHSP1 may be involved in oxidative stress via traffic between stress granules and processing bodies (12Hou H. Wang F. Zhang W. Wang D. Li X. Bartlam M. Yao X. Rao Z. J. Biol. Chem. 2011; 286: 9623-9635Abstract Full Text Full Text PDF PubMed Scopus (23) Google Scholar). Although posttranscriptional modification and the clinical relevance of CARHSP1 have been gradually recognized, there is an extremely limited understanding of the function of CARHSP1 in metabolism. Here, we demonstrate that CARHSP1 can potently inhibit the expression of gluconeogenic genes, including G6Pc and PEPCK1, when overexpressed in hepatocytes. Our data suggest that CARHSP1, via inhibition of PPARα, is of major importance in hepatic metabolism. 8- to 10-week-old male C57BL/6J mice and PPARα knockout (B6, 129S4-Pparatm1Gonz/J) mice were purchased from The Jackson Laboratory (Bar Harbor, ME). Adenovirus infections were performed by tail vein injection of 2 × 109 adenoviral particles per mouse. Blood glucose was measured by tail bleeds using an Ascencia Elite (Bayer) meter. Wild-type and db/db mice were fasted for 18 h. At time 0, blood glucose was measured, and immediately thereafter, 2 g pyruvate (dissolved in PBS)/kg body weight were injected intraperitoneally. Blood glucose was measured at indicated time points. Mice were fed a standard diet (22.5% protein, 11.8% fat, and 52% carbohydrate by mass) and sacrificed by CO2 asphyxiation 4–5 days after adenovirus injection. All animal work was performed in accordance with the University of Michigan Animal Care and Use Committee. Reagents were from the following sources. Forskolin, dexamethasone, insulin, GW6471, and an antibody against FLAG were purchased from Sigma-Aldrich (St. Louis, MO). Antibodies against CARHSP1, PPARα, β-actin, peroxisome proliferator-activated receptor γ coactivator 1-α (PGC-1α), GFP, HNF4α, and GAPDH were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). Antibodies against Myc-tag and phospho-Ser/Thr were from Cell Signaling Technology, Inc. and Spring Bioscience, respectively. Primary mouse hepatocytes and HepG2 cell lines were grown in DMEM high glucose supplemented with 10% FBS and 50 mg/ml of a Pen/Strep mix in a 37 °C/5% CO2 humidified incubator. Desired DNA fragments encoding different lengths of the G6Pc promoter region were PCR-amplified from human genomic DNA and inserted into the pGL4.11 luciferase reporter vector (Promega, Madison, WI). The inserts were positioned between KpnI and XhoI sites relative to the luciferase coding sequence. Proper insertion was verified by direct DNA sequencing. To construct a pGL4–159-Luc vector containing a mutation in the PPAR-responsive element (pGL4–159mut-Luc), site-directed mutagenesis of the pGL4–159-Luc vector was carried out using PCR methods according to the manufacturer's recommendations (Agilent Technologies). The synthetic oligonucleotide primers used for mutagenesis were 5′-CAAACGTGGTTTTTGGTTCCAACGAGCAGGGCTGGGTTGACCTG-3′ (sense) and 5′-CAGGTCAACCCAGCCCTGCTCGTTGGAACCAAAACCACGTTTG-3′ (antisense). The coding region sequences corresponding to human CARHSP1-GFP and FLAG-PPARα (full-length and different-length fragments), as well as human HFN4α, were amplified from human cDNA by high-fidelity pfu polymerase (Agilent Technologies). PCR products were sequenced and cloned into pcDNA3.1. HepG2 cells were seeded 24 h before transfection at 50–60% confluence and then cotransfected with plasmids with Lipofectamine 2000 (Invitrogen). Luciferase activity was detected with a luciferase substrate kit (Promega). Primary mouse hepatocytes were reverse-transfected with siRNA-CARHSP1 (s78866, Ambion, Inc.) using Lipofectamine RNAiMAX Reagent (Invitrogen) according to the manufacturer's recommendations. Regarding adenovirus infection, hepatocytes were infected with adenoviruses in 10% FBS DMEM for 4 h and then changed to fresh 10% FBS DMEM. To generate adenoviral vectors for overexpressing CARHSP1 or PPARα, the coding region sequences corresponding to human CARHSP1 and PPARα were amplified from human cDNA by high-fidelity pfu polymerase (Agilent Technologies). PCR products were sequenced and cloned into AdTrack-CMV from Agilent Technologies. Next, the gene coding sequences were cloned from Ad-track to the Ad-Easy vector by homologous recombination in Escherichia coli. To package the adenoviruses, adenoviral vectors were linearized with the restriction enzyme, PacI, and transfected into HEK293 cells using Lipofectamine 2000. After propagation, the recombinant adenoviruses were purified by CsCl2 density gradient ultracentrifugation. Adenovirus genomic DNA was purified with the NucleoSpin virus kit (Macherey Nagel), and adenovirus titration was performed using the Adeno-Xtm quantitative PCR titration kit (Clontech). HepG2 cells were lysed in lysis buffer (50 mm Tris-HCl (pH 7.8), 137 mm NaCl, 1 mm EDTA) containing 0.1% Triton-X-100 and a protease inhibitor mixture (Roche). After centrifugation at 12,000 rpm for 15 min at 4 °C, the supernatants were collected for an immunoprecipitation (IP) assay. Cellular extracts were precleared with protein G plus agarose for 1 h at 4 °C and then incubated with an anti-CARHSP1 or anti-PPARα (Santa Cruz Biotechnology, Inc.) antibody overnight at 4 °C. Normal IgG was used for a negative control. The immunocomplexes were pulled down by incubation with protein G-agarose for 1 h at 4 °C and washed four times with wash buffer (20 mm, 0.2 mm EDTA, 100 mm KCl, 2 mm MgCl2, 0.1% Tween 20, 10% glycerol). The samples were separated by SDS-PAGE and analyzed by immunoblotting using an anti-PPARα or -CARHSP1 antibody. Total RNA from liver samples of individual animals was extracted using TRIzol reagent (Invitrogen) according to the manufacturer's instructions. Total RNA from cells in culture was extracted using the RNeasy kit (Qiagen). RNA was reverse-transcribed into cDNA with SuperScript III (Invitrogen) and oligo-dT20 primers (Invitrogen). The abundance of transcripts was assessed by a real-time PCR system (Bio-Rad) using iQ SYBR Green Supermix (Bio-Rad). The relative quantification for each gene of interest was normalized against the internal control, 18S. The primer sequences are shown in supplemental Table 1. Primary mouse hepatocytes were isolated from 8- to 10-week-old mice as described previously (13Xu J. Xiao G. Trujillo C. Chang V. Blanco L. Joseph S.B. Bassilian S. Saad M.F. Tontonoz P. Lee W.N. Kurland I.J. J. Biol. Chem. 2002; 277: 50237-50244Abstract Full Text Full Text PDF PubMed Scopus (123) Google Scholar). In brief, mice were anesthetized and the liver was exposed. A syringe pump was set up with attached silastic tubing and then inserted into the portal vein. The liver was perfused with liver perfusion medium and liver digestion medium (Invitrogen). Hepatocytes were isolated by shaking the liver in Leibovitz L-15 medium (Invitrogen) on ice. Hepatocytes were washed twice and separated from other types of cells with Percoll (Sigma). Hepatocytes were seeded on rat tail type I collagen-coated plates or dishes (BD Biosciences) in Williams' E medium (Invitrogen) supplemented with 10% FBS for 4 h, followed by a change to fresh 10% FBS DMEM. All mammalian cell extracts were prepared with lysis buffer (Thermo Scientific) that included the following: 50 mm Tris (pH 7.5), 120 mm NaCl, 1 mm EDTA, 6 mm EGTA, 0.1% Nonidet P-40, 20 mm NaF, 1 mm sodium pyrophosphate, 30 mm 4-nitrophenyl phosphate, 1 mm benzamidine, and a protease inhibitor mixture (Roche). Cells were rinsed twice with ice-cold PBS and incubated on ice for 30 min. Cytoplasmic and nuclear extracts were prepared with NE-PER® nuclear and cytoplasmic extraction reagents (Thermo Scientific). Protein extracts were resolved by SDS-PAGE using 12% gels and electroblotted onto PVDF membranes (Bio-Rad). Membranes were blocked for 1 h at room temperature in TBS Tween 20 containing 5% (w/v) nonfat dry milk powder, washed twice, and incubated overnight with primary antibodies diluted 1:200–1000 in 5% nonfat milk solution. After further washing, membranes were incubated with a donkey anti-rabbit or mouse IRDye-conjugated IgG (Li-Cor Odyssey) secondary antibody diluted 1:5000 for 1 h. Blots were scanned, and the image was displayed in grayscale. The intensity of the protein bands was quantified using an image processing program (Li-Cor Odyssey). Mouse primary hepatocytes were infected with adenoviruses (50 m.o.i./each adenovirus) for 24 h in 10% FBS DMEM, and then maintained in 0.2% FBS DMEM for another 24 h. Next, media were replaced with Krebs-Ringer buffer (115 mm NaCl, 5.9 mm KCl, 1.2 mm MgCl2, 2.5 mm CaCl2, 25 mm NaHCO3, 1.2 mm NaH2PO4 (pH 7.4)) plus 10 mm lactate and 1 mm pyruvate. Eight hours later, the glucose released in the media was measured using a QuantiChromtm glucose assay kit (Bioassay Systems, CA). ChIP assays were performed according to the manufacturer's instructions with minor modifications using the EZ ChIP kit (Millipore). In brief, liver tissue or hepatocytes were treated for 15 min with 1% formaldehyde at room temperature for cross-linking, and these reactions were terminated by the addition of glycine at a final concentration of 125 mm. Liver tissue or cells were lysed and chromatin extracts were sonicated for obtaining DNA fragments between 500–1000 bp. The sonicated chromatin was first precleared for 1 h with protein G-agarose. After centrifugation, supernatants were incubated overnight at 4 °C with 5 μg anti-PPARα antibody (Santa Cruz Biotechnology, Inc.) or normal rabbit IgG. The immunoprecipitated DNA-protein complex was incubated with protein G-agarose for 1 h at 4 °C. After centrifugation, the complex was washed in low-salt buffer, high-salt buffer, LiCl buffer, and Tris-EDTA buffer. The protein-chromatin cross-linking in the immunoprecipitated complex was reversed at 65 °C overnight. Proteins were eliminated using Proteinase K for 30 min at 45 °C. DNA was purified and used as a template for real-time PCR. The PCR primers used for the analysis of G6Pc and PEPCK1 promoters are listed in supplemental Table 1. PCR amplification products were confirmed on ethidium bromide-stained 2% agarose gels. Statistical comparisons and analyses between two groups were performed by two-tailed unpaired Student's t test, and among three groups or more they were performed by one-way analysis of variance followed by a Newman-Keuls test. A p value of < 0.05 was considered statistically significant. Data are presented as mean ± S.E. To determine whether CARHSP1 is a potential regulator of metabolism, we first detected its expression in the liver. CARHSP1 protein levels were significantly reduced after mice were fasted for 24 h. Furthermore, CARHSP1 protein expression was increased after mice were refed for 1 h and returned to base line after 6 h of refeeding (Fig. 1, A and B), although CARHSP1 mRNA expression levels remained low (C). To mimic fasting signals in vivo, primary mouse hepatocytes were isolated and incubated in vitro with forskolin (FSK) and dexamethasone (DEX) (14Rodgers J.T. Haas W. Gygi S.P. Puigserver P. Cell Metab. 2010; 11: 23-34Abstract Full Text Full Text PDF PubMed Scopus (95) Google Scholar) in a low-nutrient status (0.2% FBS medium) for 16 h. CARHSP1 protein levels were significantly reduced in these altered culture conditions (Fig. 1D). CARHSP1 mRNA expression was down-regulated, whereas G6Pc and PEPCK1 (phosphoenolpyruvate carboxykinase 1, soluble) mRNA levels were up-regulated by stimulation with forskolin and dexamethasone (Fig. 1E). G6Pc and PEPCK1 are regulated at the transcriptional level by a variety of hormonal and nutrient signals including insulin, glucocorticoids, thyroid hormones, and cyclic AMP (15Hall R.K. Granner D.K. J. Basic Clin. Physiol. Pharmacol. 1999; 10: 119-133Crossref PubMed Scopus (47) Google Scholar, 16Hanson R.W. Reshef L. Annu. Rev. Biochem. 1997; 66: 581-611Crossref PubMed Scopus (634) Google Scholar). Our data suggest that fasting conditions contribute to the changes of CARHSP1 expression in the liver. Insulin signaling is activated during refeeding. We found that insulin increases CARHSP1 expression but decreases G6Pc expression in primary mouse hepatocytes (Fig. 2, A–C). Insulin treatment also induced CARHSP1 phosphorylation, whereas wortmannin, a PI3K inhibitor, blocked phosphorylation (Fig. 2D). Regarding the subcellular protein location of CARHSP1, we demonstrated that CARHSP1 is located in both the cytoplasm and nucleus of hepatocytes (Fig. 2E). Furthermore, we found that insulin induced CARHSP1 nuclear translocation in hepatocytes (Fig. 2F). All in all, our data suggest that regulation of CARHSP1 expression and subcellular localization occur during fasting and refeeding conditions.FIGURE 2Subcellular distribution of CARHSP1 in hepatocytes. A, primary mouse hepatocytes were treated with insulin (50 nm) in 2% FBS DMEM and the expression levels of CARHSP1 (A) and of G6Pc (B) were determined by real-time PCR at different time points. C, primary mouse hepatocytes were treated with insulin at different dosages for 24 h, and the expression of CARHSP1was determined by real-time PCR. D, HepG2 cells were incubated with insulin (50 nm) for the indicated times. Cells treated with insulin for 30 min were pretreated with wortmannin (200 nm). Cell extracts were subjected to coimmunoprecipitation with an antibody against CARHSP1 and then immunoblotting with an antibody against phosphorylated Ser/Thr (pan). E, cytoplasmic and nuclear aliquots (20 μg loaded protein) were purified from HepG2 cells and subjected to immunoblotting analysis. F, primary mouse hepatocytes were treated with insulin (50 nm) for different times. Cytoplasmic and nuclear protein were extracted and subjected to immunoblotting analysis. Nuclear CARHSP1 is quantitatively analyzed and normalized against Lamin A/C. Data are presented as mean ± S.E. *, p < 0.05; **p < 0.01.View Large Image Figure ViewerDownload Hi-res image Download (PPT) To determine the function of CARHSP1 in hepatic metabolism, we injected Ad-CARHSP1 into C57BL/6J mice by tail vein to obtain CARHSP1 gain of function. The expression of Ad-CARHSP1 in the liver was confirmed by Western blotting (Fig. 3A). Interestingly, CARHSP1 gain of function in the liver significantly inhibited gluconeogenic gene expression at 4 days after adenovirus tail vein injection (Fig. 3B). Moreover, in cultured primary mouse hepatocytes, CARHSP1 dramatically inhibited G6Pc and PEPCK1 expression both in low serum (0.2% serum) and upon further induction in the presence of forskolin (10 μm) and dexamethasone (1 μm) (Fig. 3C). A similar result was observed in the CARHSP1-overexpressing HepG2 human hepatocyte cell line (Fig. 3D). CARHSP1 also decreased the protein expression of G6Pc and PEPCK1 in primary mouse hepatocytes (Fig. 3E). Functionally, CARHSP1 overexpression potently inhibited glucose output from primary mouse hepatocytes (Fig. 3F). Hepatic glucose output generally reflects the total flux resulting from gluconeogenic and glycogenolytic pathways (17Vega R.B. Huss J.M. Kelly D.P. Mol. Cell. Biol. 2000; 20: 1868-1876Crossref PubMed Scopus (952) Google Scholar). Overall, our data suggest that CARHSP1 inhibits glucose output at least partly through modulation of gluconeogenesis. To determine whether CARHSP1 functions in a physiological context, we investigated the effect of endogenous CARHSP1 on the expression of gluconeogenic genes. Using siRNA against CARHSP1, the expression of CARHSP1 was efficiently knocked down in hepatocytes at both the protein and mRNA levels (Fig. 4, A and B). Consistent with our observation that CARHSP1 gain of function in hepatocytes resulted in marked down-regulation of gluconeogenic genes, CARHSP1 loss of function, conversely, increased the expression of G6Pc and PEPCK1 (Fig. 4, C and D) both in basal and forskolin+DEX-stimulated conditions. This indicates that loss of CARHSP1 is probably releasing basal inhibition of these genes. CARHSP1 knockdown also significantly increases glucose output from primary mouse hepatocytes (Fig. 4E). Thus, CARHSP1 is required to maintain steady basal levels of expression and ensure homeostatic regulation of gluconeogenic genes. Previous studies identified that PPARα could regulate gluconeogenic genes in both wild-type (18Im S.S. Kim M.Y. Kwon S.K. Kim T.H. Bae J.S. Kim H. Kim K.S. Oh G.T. Ahn Y.H. J. Biol. Chem. 2011; 286: 1157-1164Abstract Full Text Full Text PDF PubMed Scopus (44) Google Scholar) and DEX-treated LDL receptor (LDL-R) null mice (19Bernal-Mizrachi C. Weng S. Feng C. Finck B.N. Knutsen R.H. Leone T.C. Coleman T. Mecham R.P. Kelly D.P. Semenkovich C.F. Nat. Med. 2003; 9: 1069-1075Crossref PubMed Scopus (177) Google Scholar). We found that CARHSP1 inhibits PPARα-induced expression of G6Pc and PEPCK1 by as much as 80% in HepG2 cells (Fig. 5A) and potently inhibits PPARα-induced glucose output in primary hepatocytes (B). Therefore, our data suggest that there is an inhibitory epistatic effect of CARHSP1 on PPARα activity in the regulation of hepatic gluconeogenesis. Next, we sought to investigate the mechanisms linking CARHSP1 and PPARα in hepatic gluconeogenesis. CARHSP1 could impair PPARα activity in different ways (i.e. affecting protein levels, subcellular protein localization, or DNA binding). However, we found that CARHSP1 did not alter PPARα protein expression in HepG2 cells (Fig. 5C). As determined by a coimmunoprecipitation assay, endogenous CARHSP1 physically interacts with PPARα in vivo (Fig. 5D) and in vitro (E). To determine the respective binding domains of both CARHSP1 and PPARα, we generated CARHSP1 fragment-GFP fusion proteins and FLAG-PPARα fragments to perform immunoprecipitation assays. We found the 1–60-amino acid (aa) sequence of CARHSP1 binds to PPARα (Fig. 6A), whereas the 167–244-aa sequence of PPARα binds to CARHSP1, respectively (Fig. 6B). Interestingly, the PPARα agonist WY-14643 can induce the dissociation of CARHSP1 from PPARα (Fig. 6C). To determine whether CARHSP1 competes with coactivators of PPARα, we performed coimmunoprecipitation and ChIP assays. PGC-1α is a well known PPARα coactivator that plays a critical role in gluconeogenesis. We demonstrated that CARHSP1 reduced the binding of PGC-1α to PPARα in hepatocytes (Fig. 6D). Moreover, CARHSP1 could suppress the binding of PGC-1α to the G6Pc promoter, where there exists a PPRE site for PPARα binding (Fig. 6E). To further establish whether CARHSP1 regulates gluconeogenic gene expression at the transcriptional level, we constructed pGL4 vectors with inserted DNA fragments in varying lengths encoding the G6Pc promoter. Overexpression of PPARα resulted in increased activation of the G6Pc promoter (-1035/+80, −520/+80, and −159/+80) in HepG2 cells, which can be readily inhibited by CARHSP1 (Fig. 7A). Furthermore, it should be noted that CARHSP1 can inhibit G6Pc promoter activity in the absence of overexpressed PPARα, consistent with its ability to regulate basal levels of G6Pc in these cells, as described above (Fig. 3). However, PPARα cannot activate the −52/+80 fragment of the G6Pc promoter, which implies that a functional binding site may exist between −159 bp and −52 bp in the G6Pc promoter. Analysis of transcriptional binding sites with Genomatix software showed that highly conserved PPAR-response elements (PPREs) exist in both human and mouse G6Pc promoters. The putative PPRE (-75/-63) was mutated to determine whether this PPRE is a functional PPARα binding element. The G6Pc-PPREmut promoter could not be activated by PPARα. Furthermore, CARHSP1 had no inhibitory effect on the activity of the mutated promoter (Fig. 7B), suggesting CARHSP1-mediated down-regulation of the G6Pc gene via PPARα. Next, CHIP assays were performed to further determine the effect of CARHSP1 on PPARα binding to the promoter regions of gluconeogenic genes. Consistent with the data shown above, we demonstrate that PPARα can directly bind to the putative PPRE (-75/-63 bp) in the G6Pc promoter and that CARHSP1 potently inhibits the binding of PPARα to this PPRE in human hepatocytes infected with Ad-CARHSP1 (Fig. 7C). Similar results were observed regarding CARHSP1 inhibition of PPARα binding to the PPRE (-387/-399) existing in the human PEPCK1 promoter in HepG2 cells (Fig. 7C). Our data also demonstrate that CARHSP1 has a potent inhibitory effect on the binding of PPARα to G6Pc and PEPCK1 promoters in the mouse liver 4 days after delivering Ad-CARHSP1 by the tail vein (Fig. 7D). Thus, our data identified that CARHSP1 inhibits G6Pc and PEPCK1 expression at the transcriptional level through inhibition of PPARα activity. To determine the repressive domain of CARHSP1, we generated different lengths of the CARHSP1 coding region. The N-terminal 31–65-amino acid sequence of CARHSP1 is necessary for CARHSP1 to fulfill its inhibitory effect on PPARα-induced activation of G6Pc promoter (Fig. 7, E and F). Next, we determined whether CARHSP1 interacts with other nuclear transcriptional factors. HNF4α is a well demonstrated transcriptional factor in regulating gluconeogenesis. However, we did not observe an interaction between CARHSP1 and HNF4α (Fig. 8A), and CARHSP1 did not significantly inhibit HNF4α-induced expression of G6Pc and PEPCK1 (B). To determine whether PPARα is an essential mediator of CARHSP1-induced inhibition of gluconeogenesis, we pharmacologically inactivated PPARα through administration of the PPARα antagonist GW6471, which induces a PPARα conformational change followed by the recruitment of corepressors (20Xu H.E. Stanley T.B. Montana V.G. Lambert M.H. Shearer B.G. Cobb J.E. McKee D.D. Galardi C.M. Plunket K.D. Nolte R.T. Parks D.J. Moore J.T. Kliewer S.A. Willson T.M. Stimmel J.B. Nature. 2002; 415: 813-817Crossref PubMed Scopus (520) Google Scholar). Interestingly, after GW6471 treatment (20 μm), G6Pc and PEPCK1 expression levels did not change when HepG2 cells were treated with Ad-CARHSP1, which indicated that the regulation of gluconeogenic genes by CARHSP1 was blocked dramatically when PPARα was antagonized (Fig. 8, C and D). We further examined the necessary role of PPARα in mediating the effect of CARHSP1 on gluconeogenesis in hepatocytes isolated from PPARα knockout mice. Consistent with what we observed in pharmacologically PPARα-inactivated hepatocytes, we demonstrated that CARHSP1 also lost its inhibitory effect on G6Pc and PEPCK1 expression in PPARα knockout hepatocytes (Fig. 8, E and F). These data support our hypothesis that CARHSP1, by inhibiting PPARα activity, is a negative regulator of hepatic gluconeogenic genes. Because CARHSP1 overexpression results in down-regulation of gluconeogenic genes, we sought to test the hypothesis that increases in CARHSP1 will suppress hepatic gluconeogenesis in vivo. As shown in Fig. 9A, overexpression of CARHSP1 reduced fasting blood glucose levels in C57BL/6J mice. More interestingly, the pyruvate sodium tolerance tests performed in our study demonstrate that changes in blood glucose levels were significantly reduced in CARHSP1-treated a" @default.
• W2093961921 created "2016-06-24" @default.
• W2093961921 creator A5046229622 @default.
• W2093961921 creator A5054309140 @default.
• W2093961921 creator A5079226908 @default.
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• W2093961921 date "2011-11-01" @default.
• W2093961921 modified "2023-09-27" @default.
• W2093961921 title "Inhibition of Gluconeogenic Genes by Calcium-regulated Heat-stable Protein 1 via Repression of Peroxisome Proliferator-activated Receptor α" @default.
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