Inhibition of Gluconeogenic Genes by Calcium-regulated Heat-stable Protein 1 via Repression of Peroxisome Proliferator-activated Receptor α*

Background: Gluconeogenesis contributes to insulin resistance in type 1 and type 2 diabetes, but underlying molecular mechanisms remain unclear. Results: CARHSP1 functions at the transcriptional level to negatively regulate gluconeogenic genes in the liver. Conclusion: CARHSP1 inhibits hepatic gluconeogenic gene expression via repression of PPARα. Significance: CARHSP1 is a negative regulator of hepatic gluconeogenesis and a potential 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 (1)(2)(3) with increased gluconeogenesis as a major contributor to fasting hyperglycemia in both type 1 and type 2 diabetes (4,5). Gluconeogenesis is controlled by certain rate-limiting enzymes such as the glucose-6-phosphatase catalytic subunit (G6Pc) 3 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.
Calcium-regulated heat-stable protein (CARHSP1) is a ubiquitously expressed phosphoprotein that is comprised of 147 amino acids with nearly 14% proline (6). CARHSP1 is serinephosphorylated 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) (7). On the other hand, CARHSP1 can be dephosphorylated by Ca 2ϩ /calmodulin-regulated protein phosphatase calcineurin (PP2B) (8). Coordinated phosphorylation and dephosphorylation of CARHSP1 contribute to the multiple phosphorylated isoforms of CARHSP1 in acinar cells (9). Recently, CARHSP1 was discovered as a biomarker for diabetic retinopathy (10) and as a regulator of TNF-␣ mRNA stability in macrophages (11). CARHSP1 may be involved in oxidative stress via traffic between stress granules and processing bodies (12). 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 CAR-HSP1 can potently inhibit the expression of gluconeogenic genes, including G6Pc and PEPCK1, when overexpressed in hepatocytes. Our data suggest that CARHSP1, via inhibition of PPAR␣, is of major importance in hepatic metabolism.

EXPERIMENTAL PROCEDURES
Animal Procedures-8-to 10-week-old male C57BL/6J mice and PPAR␣ knockout (B6, 129S4-Ppara tm1Gonz /J) mice were purchased from The Jackson Laboratory (Bar Harbor, ME). Adenovirus infections were performed by tail vein injection of 2 ϫ 10 9 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 CO 2 asphyxiation 4 -5 days after adenovirus injection. All animal work was performed in accordance with the University of Michigan Animal Care and Use Committee.
Cell Culture-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% CO 2 humidified incubator.
Construction of Plasmids and Transfections-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 PPARresponsive 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 5Ј-CAAACGTGGTTTTT-GGTTCCAACGAGCAGGGCTGGGTTGACCTG-3Ј (sense) and 5Ј-CAGGTCAACCCAGCCCTGCTCGTTGGAACCAA-AACCACGTTTG-3Ј (antisense). The coding region sequences corresponding to human CARHSP1-GFP and FLAG-PPAR␣ (full-length and different-length fragments), as well as human HFN4␣, 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).
Construction of Adenoviruses-To generate adenoviral vectors for overexpressing CARHSP1 or PPAR␣, the coding region sequences corresponding to human CARHSP1 and PPAR␣ 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 CsCl 2 density gradient ultracentrifugation. Adenovirus genomic DNA was purified with the NucleoSpin virus kit (Macherey Nagel), and adenovirus titration was performed using the Adeno-X TM quantitative PCR titration kit (Clontech).
Coimmunoprecipitation-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-CAR-HSP1 or anti-PPAR␣ (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 MgCl 2 , 0.1% Tween 20, 10% glycerol). The samples were separated by SDS-PAGE and analyzed by immunoblotting using an anti-PPAR␣ or -CARHSP1 antibody.
Total RNA Preparation and RT Quantitative PCR Analysis-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 reversetranscribed 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.
Isolation of Primary Mouse Hepatocytes-Primary mouse hepatocytes were isolated from 8-to 10-week-old mice as described previously (13). 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.
Cell Extraction and Western Blotting-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).
ChIP-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 g anti-PPAR␣ 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 Analysis-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 Ͻ 0.05 was considered statistically significant. Data are presented as mean Ϯ S.E.

RESULTS
The Expression of CARHSP1 Changes with Nutrient Transition-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) (14) 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 (15,16). 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.
Hepatic Gluconeogenic Genes Are Down-regulated by CARHSP1-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 M) and dexamethasone (1 M) (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 (17). Overall, our data suggest that CAR-HSP1 inhibits glucose output at least partly through modulation of gluconeogenesis.
CARHSP1 Knockdown Up-regulates the Expression of Gluconeogenic Genes-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  . 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 g 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 Ϯ S.E. *, p Ͻ 0.05; ** p Ͻ 0.01. NOVEMBER 25, 2011 • VOLUME 286 • NUMBER 47

JOURNAL OF BIOLOGICAL CHEMISTRY 40587
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 forskolinϩDEX-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.

CARHSP1 Inhibits PPAR␣-induced Gluconeogenic Gene
Expression in Hepatocytes-Previous studies identified that PPAR␣ could regulate gluconeogenic genes in both wild-type (18) and DEX-treated LDL receptor (LDL-R) null mice (19). We found that CARHSP1 inhibits PPAR␣-induced expression of G6Pc and PEPCK1 by as much as 80% in HepG2 cells (Fig. 5A) and potently inhibits PPAR␣-induced glucose output in primary hepatocytes (B). Therefore, our data suggest that there is an inhibitory epistatic effect of CARHSP1 on PPAR␣ activity in the regulation of hepatic gluconeogenesis. Next, we sought to investigate the mechanisms linking CARHSP1 and PPAR␣ in hepatic gluconeogenesis. CARHSP1 could impair PPAR␣ activity in different ways (i.e. affecting protein levels, subcellular protein localization, or DNA binding). However, we found that CARHSP1 did not alter PPAR␣ protein expression in HepG2 cells (Fig. 5C). As determined by a coimmunoprecipitation assay, endogenous CARHSP1 physically interacts with PPAR␣ in vivo (Fig. 5D) and in vitro (E).
CARHSP1 Interacts with PPAR␣ in Hepatocytes-To determine the respective binding domains of both CARHSP1 and PPAR␣, we generated CARHSP1 fragment-GFP fusion proteins and FLAG-PPAR␣ fragments to perform immunoprecipitation assays. We found the 1-60-amino acid (aa) sequence of CARHSP1 binds to PPAR␣ (Fig. 6A), whereas the 167-244-aa sequence of PPAR␣ binds to CARHSP1, respectively (Fig. 6B). Interestingly, the PPAR␣ agonist WY-14643 can induce the dissociation of CARHSP1 from PPAR␣ (Fig. 6C). To determine whether CARHSP1 competes with coactivators of PPAR␣, we performed coimmunoprecipitation and ChIP assays. PGC-1␣ is a well known PPAR␣ coactivator that plays a critical role in gluconeogenesis. We demonstrated that CARHSP1 reduced the binding of PGC-1␣ to PPAR␣ in hepatocytes (Fig. 6D). Moreover, CARHSP1 could suppress the binding of PGC-1␣ to the G6Pc promoter, where there exists a PPRE site for PPAR␣ binding (Fig. 6E).
CARHSP1 Inhibits the Binding of PPAR␣ to the G6Pc Promoter-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 PPAR␣ resulted in increased activation of the G6Pc promoter (-1035/ϩ80, Ϫ520/ϩ80, and Ϫ159/ϩ80) 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 PPAR␣, consistent with its ability to regulate basal levels of G6Pc in these cells, as described above (Fig. 3). However, PPAR␣ cannot activate the Ϫ52/ϩ80 fragment of the G6Pc promoter, which implies that a functional binding site may exist between Ϫ159 bp and Ϫ52 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 PPAR␣ binding element. The G6Pc-PPREmut promoter could not be activated by PPAR␣. 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 PPAR␣.
Next, CHIP assays were performed to further determine the effect of CARHSP1 on PPAR␣ binding to the promoter regions of gluconeogenic genes. Consistent with the data shown above, we demonstrate that PPAR␣ can directly bind to the putative PPRE (-75/-63 bp) in the G6Pc promoter and that CARHSP1 potently inhibits the binding of PPAR␣ to this PPRE in human hepatocytes infected with Ad-CARHSP1 (Fig. 7C). Similar results were observed regarding CARHSP1 inhibition of PPAR␣ 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 PPAR␣ 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 PPAR␣ 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 PPAR␣-induced activation of G6Pc promoter (Fig. 7, E and F).
PPAR␣ Is Required for CARHSP1 Inhibition of Gluconeogenic Gene Expression-Next, we determined whether CARHSP1 interacts with other nuclear transcriptional factors. HNF4␣ is a well demonstrated transcriptional factor in regulating gluconeogenesis. However, we did not observe an interaction  between CARHSP1 and HNF4␣ (Fig. 8A), and CARHSP1 did not significantly inhibit HNF4␣-induced expression of G6Pc and PEPCK1 (B). To determine whether PPAR␣ is an essential mediator of CARHSP1-induced inhibition of gluconeogenesis, we pharmacologically inactivated PPAR␣ through administration of the PPAR␣ antagonist GW6471, which induces a PPAR␣ conformational change followed by the recruitment of corepressors (20). Interestingly, after GW6471 treatment (20 M), 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 PPAR␣ was antagonized (Fig. 8, C  and D). We further examined the necessary role of PPAR␣ in mediating the effect of CARHSP1 on gluconeogenesis in hepatocytes isolated from PPAR␣ knockout mice. Consistent with what we observed in pharmacologically PPAR␣-inactivated hepatocytes, we demonstrated that CARHSP1 also lost its inhibitory effect on G6Pc and PEPCK1 expression in PPAR␣ knockout hepatocytes (Fig. 8, E and F). These data support our hypothesis that CARHSP1, by inhibiting PPAR␣ activity, is a negative regulator of hepatic gluconeogenic genes.
CARHSP1 Suppresses Hepatic Gluconeogenesis in Vivo-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 animals at the specified time points (15,75, and 90 min) (Fig. 9B). All in all, our data suggest that CARHSP1 potently inhibits hepatic gluconeogenesis and contributes to glucose homeostasis.

DISCUSSION
In this study, we demonstrate for the first time that CAR-HSP1 has a novel and potent inhibitory effect on hepatic gluconeogenic gene expression and may contribute to the improvement of insulin resistance in diabetic mice. Here, another novel finding is that CARHSP1 fulfills its function through inhibition of PPAR␣ activity.
CARHSP1 is a cold shock domain (CSD)-containing protein with a high level of resemblance to cold shock proteins (6). All living organisms must adapt to environmental changes, including cold shock, heat shock, and nutritional status. Importantly, the nutritional status of the extracellular environment must be sensed and information needs to be conducted to intracellular signaling pathways by cells themselves. Cold shock proteins in FIGURE 6. CARHSP1 interacts with PPAR␣. A, either full-length CARHSP1-GFP or the 1-60 amino acid sequence of the CARHSP1-GFP fusion protein was cotransfected with FLAG-PPAR␣ into HepG2 cells for 24 h before coimmunoprecipitation with an antibody against FLAG and then immunoblotting with an antibody against GFP. B, FLAG-PPAR␣ fragments and Myc-tagged CARHSP1 were cotransfected into HepG2 cells for 24 h before coimmunoprecipitation with an antibody against Myc-tag and then by immunoblotting (IB) with an antibody against FLAG. C, Primary mouse hepatocytes were treated with WY-14643 (50 M) for 40 min and then subjected to immunoprecipitation with an antibody against PPAR␣. Normal rabbit IgG was used as a negative control. D, HepG2 cells were infected with Ad-CARHSP1 or Ad-LacZ 24 h before coimmunoprecipitation with an antibody against PPAR␣ and then subjected to immunoprecipitation with an antibody against PGC1␣ or PPAR␣. E, C57BL/6J mice were injected with Ad-CARHSP1 or Ad-LacZ (2 ϫ 10 9 virus particles) by tail vein. After 4 days, liver samples were isolated, and then CHIP assays were performed using antibodies against PGC-1␣ and normal rabbit IgG. Purified DNA fragments were detected by real-time PCR. The primers for the ChIP assays were designed to detect the DNA sequence containing putative PPREs in the promoter of G6Pc. Data shown are presented as mean Ϯ S.E. *, p Ͻ 0.05.
both prokaryotes and eukaryotes are susceptible to environmental changes, especially abrupt drops in temperature (21)(22)(23). These DNA-and RNA-binding proteins are often considered to be transcriptional and/or translational regulatory proteins (24).
In the biological context, a complicated regulatory network consisting of numerous transcription factors and coregulators controls gluconeogenesis (25). Transcription factors or coregulators such as PGC-1␣ (26), forkhead box protein O1 (FOXO1) (27,28), cAMP response element-binding protein-regulated transcription coactivator 2 (CRTC2) (29,30) and HNF4 (31) can adapt to cellular nutrient status, as shown by changes in the expression of these proteins. The up-regulation or down-regulation of these transcription factors/coregulators has been shown to be important for normal cellular metabolism and function. Here, we identified that CARHSP1, as a stress-sensitive gene, is highly relevant to glucose metabolism. CARHSP1 protein expression is down-regulated after fasting and then returns to base line after refeeding for 6 h, which is not consistent with mRNA. To our knowledge, this may be due to increased protein stability and increased translation of CAR-HSP1 protein after refeeding. As a phosphoprotein, CARHSP1 is phosphorylated at multiple sites (7)(8)(9). Our data suggest that the 31-65 amino acid sequence located in the N-terminal domain, which is rich in phosphorylation sites, is necessary for CARHSP1 to inhibit PPAR␣ activity (Fig. 7F). Although a transgenic CARHSP1 mouse line would be a much better model to study the exact role of CARHSP1 in an altered nutrient status, it FIGURE 7. CARHSP1 inhibits PPAR␣-mediated transcriptional regulation of gluconeogenic genes. A, the effect of CARHSP1 on PPAR␣-induced activation of the G6Pc promoter. Serial deletion constructs of the G6Pc promoter were cotransfected with TK-RL into HepG2 cells. After 12 h, cells were infected with Ad-LacZ or Ad-CARHSP1 in the presence or absence of Ad-PPAR␣ for another 48 h. Promoter activities of serial deletion constructs were detected and normalized to Renilla activity. B, CARHSP1 had no effect on the activity of the G6Pc promoter (-159/ϩ80) in which the PPRE was mutated. HepG2 cells were transfected with G6Pc promoter-luc (-159/ϩ80) or G6Pc (-159/ϩ80) Mut-luc. After 12 h, cells were infected with Ad-LacZ or Ad-CARHSP1 in the presence or absence of Ad-PPAR␣ for another 48 h. Activity of the mutated G6Pc promoter was detected and normalized to Renilla activity. The numbers indicate the distance in nucleotides from the transcription start site (ϩ1) of the human G6Pc gene. Representative data are shown from three independent experiments. Data are presented as mean Ϯ S.E. *, p Ͻ 0.05; **, p Ͻ 0.01. C, HepG2 cells were infected with Ad-LacZ or Ad-CARHSP1 (100 m.o.i.) and after 48 h, ChIP assays were performed using antibodies against PPAR␣ and normal rabbit IgG. Data shown are from three independent experiments. D, C57BL/6J mice were injected with Ad-CARHSP1 or Ad-LacZ (2 ϫ 10 9 virus particles) by tail vein, n ϭ 4. After 4 days, liver samples were isolated and then CHIP assays were performed using antibodies against PPAR␣ and normal rabbit IgG. Purified DNA fragments were detected by real-time PCR. The primers for the CHIP assays were designed to detect the DNA sequences containing putative PPREs in the promoters of the target genes. Putative PPREs are indicated in horizontal boxes (C and D). Quantitative analysis was performed and normalized against 10% Input. E and F, CARHSP1 fragments (E) and G6Pc promoter (-520)-Luc were cotransfected into HepG2 cells. After 12 h, cells were infected with Ad-LacZ or Ad-PPAR␣ in the presence or absence of Ad-PPAR␣ for another 24 h. Promoter activity was detected and normalized to Renilla activity (F). Data shown are presented as mean Ϯ S.E. *, p Ͻ 0.05; **, p Ͻ 0.01. NOVEMBER 25, 2011 • VOLUME 286 • NUMBER 47 is through acute agonism of CARHSP1 in this study that we investigated and demonstrated that CARHSP1 negatively regulates gluconeogenic gene expression.

CARHSP1 Modulates Gluconeogenic Gene Expression
The exact molecular mechanisms underlying the effects of CARHSP1 on hepatic glucose metabolism may be complicated. Here, we focused on gluconeogenesis and found that CARHSP1 suppresses the expression of gluconeogenic enzymes, including G6Pc and PEPCK1. These gluconeogenic enzymes are regulated at the transcriptional level by numerous gluconeogenesis-related transcription factors and coregulators (32)(33)(34). PPAR␣ possesses properties and functions somewhat differently from the other two PPAR subtypes, PPAR␦ and PPAR␥ (35,36). It is well known that PPAR␣ has a central role in lipid oxidation (37,38) and ketogenesis (39) in the liver. In addition, recent studies have revealed a link between PPAR␣ and hepatic gluconeogenesis (18,19,40,41). PPAR␣-deficient mice are hypoglycemic after fasting (42) and protected against high-fat diet-induced insulin resistance (43,44). In addition, PPAR␣ knockout mice show much higher levels of plasma free fatty acids compared with those of wild-type mice in the fasting state (45,46). PPAR␣ can modulate the synthesis of glucose from non-glucose substances through regulation of genes, including glycerol 3-phosphate dehydrogenase (GPDH) and glycerol kinase (41). However, little is known about whether PPAR␣ directly regulates the expression of G6Pc and PEPCK1 in hepatocytes. We demonstrate that PPAR␣ can bind to putative PPREs in both G6Pc and PEPCK1 promoters in human and mouse hepatocytes. Recently, Im et al. (18) reported results identical to ours by also showing that hepatic G6Pc is a PPAR␣ target gene.
Although the role of PPAR␣ in the liver is well dissected, the regulation of PPAR␣ function remains unclear. In this study, we propose a hypothesis that in gluconeogenesis, CARHSP1 can modulate PPAR␣ transcriptional activity and eventually inter-  . CARHSP1 suppresses hepatic gluconeogenesis. C57BL/6J mice (male, 8 -10 weeks old) were injected with Ad-CARHSP1 or Ad-LacZ (2 ϫ 10 9 virus particles) by tail vein. Four days after adenoviral injection, mice were fasted for 18 h (A and B). A, fasting glucose levels were decreased in Ad-CAR-HSP1-injected mice. B, pyruvate sodium (2 g/kg intraperitoneally) tolerance tests show that increases in blood glucose levels were attenuated in Ad-CAR-HSP1-injected mice. Blood glucose levels were determined at the indicated time points (n ϭ 8). Ⅺ, LacZ; f, CARHSP1. Data shown are presented as mean Ϯ S.E. *, p Ͻ 0.05; **, p Ͻ 0.01. fere with its function. Actually, some coregulators have been identified to activate or suppress PPAR transcriptional activity, and these coregulators fulfill their function with tissue-or cellspecific properties (32,33,38). We demonstrate that CARHSP1 acts as a PPAR␣ repressor, resulting in a dramatic reduction of PPAR␣ transcriptional activity in hepatocytes. CARHSP1 binds to the 167-244 aa sequence of PPAR␣ which is located in the PPAR␣ hinge domain. Both the coactivator, PGC-1␣, and the corepressor, N-CoR, can also bind to this particular PPAR␣ domain (17,47). Notably, CARHSP1 down-regulation of gluconeogenic genes is dependent on PPAR␣. Here, our study provides a link between CARHSP1 and the metabolic regulator, PPAR␣ as well as providing further insight into the PPAR␣ signaling network. PPAR␣ has been recently reported to be up-regulated in db/db mice (18). Our findings that CARHSP1 is a repressor of PPAR␣ and a regulator of hepatic gluconeogenic genes suggest a functional cooperation between CARHSP1 and PPAR␣ in ameliorating insulin resistance in diabetic mice.
However, we must acknowledge that CARHSP1, in addition to its functional interaction with PPAR␣, may cross-talk with other nuclear transcription factors, such as FOXO1, cAMP response element-binding protein, and the glucocorticoid receptor. To further characterize the specificity of CARHSP1, we determined whether HNF4␣ is a CARHSP1-interacting transcription factor because HNF4␣ is known to be involved in gluconeogenesis. In this study, we demonstrate that CARHSP1 does not interact with HNF4␣ (Fig. 8). Future studies will address the potential for CARHSP1 to interact with other transcription factors. Nevertheless, our data suggest that CAR-HSP1, at the very least, is a specific repressor of PPAR␣.
Currently, the mechanisms underlying the regulation of CARHSP1 in the liver are under investigation. CARHSP1 is a substrate for activated Akt and other kinases in HEK293 cells (7), and we confirm that CARHSP1 can be phosphorylated by insulin. However, other modifications of CARHSP1 in the context of hepatic metabolism must be further studied in ensuing investigations to fully understand its function in the liver. Although gain of function models were used in this study to examine the function of CARHSP1 in hepatic gluconeogenesis, liver-selective CARHSP1 knockout mice will be essential in providing additional information regarding hepatic pathophysiological processes and molecular mechanisms. In conclusion, we show that CARHSP1 is a negative regulator of hepatic gluconeogenesis and an emerging novel therapeutic target relevant for the treatment of metabolic diseases.