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Protects against Obesity-Induced Hepatic InflammationNutrition, Metabolism and Genomics Group (R.S., S.M., D.P., S.K., M.M.), Division of Human Nutrition, Wageningen University, 6700 EV Wageningen, The Netherlands; Nutrigenomics Consortium (R.S., S.K., M.M.), Wageningen Center of Food Sciences, 6709 PA Wageningen, The Netherlands; and Department of Pathology (C.M.), Radboud University Nijmegen Medical Center, 6500 HB Nijmegen, The Netherlands
Address all correspondence and requests for reprints to: Michael Müller, Ph.D., Division of Human Nutrition, Wageningen University, P.O. Box 8129, 6700 EV Wageningen, The Netherlands. E-mail: michael.muller{at}wur.nl.
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Abstract |
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(PPAR) has been shown to have a strong antiinflammatory action in liver. However, the role of PPAR in obesity-induced inflammation is much less clear. Therefore, the aim of our study was to determine whether PPAR plays a role in obesity-induced hepatic inflammation. To induce obesity, wild-type sv129 and PPAR–/– mice were exposed to a chronic high-fat diet (HFD), using a low-fat diet (LFD) as control. In wild-type mice, HFD significantly increased the hepatic and adipose expression of numerous genes involved in inflammation. Importantly, this effect was amplified in PPAR–/– mice, suggesting an antiinflammatory role of PPAR in liver and adipose tissue. Further analysis identified specific chemokines and macrophage markers, including monocyte chemotactic protein 1 and F4/80+, that were elevated in liver and adipose tissue of PPAR–/– mice, indicating increased inflammatory cell recruitment in the knockout animals. When all groups of mice were analyzed together, a significant correlation between hepatic triglycerides and expression of inflammatory markers was observed. Many inflammatory genes that were up-regulated in PPAR–/– livers by HFD were down-regulated by treatment with the PPAR ligand Wy-14643 under normal nonsteatotic conditions, either in vivo or in vitro, suggesting an antiinflammatory effect of PPAR that is independent of reduction in liver triglycerides. In conclusion, our results suggest that PPAR protects against obesity-induced chronic inflammation in liver by reducing hepatic steatosis, by direct down-regulation of inflammatory genes, and by attenuating inflammation in adipose tissue. |
Introduction |
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, -ß/
, and -
, which share a common molecular structure and mechanism of action (1). PPARs are ligand-activated transcription factors that regulate gene transcription by binding to specific DNA sequences, known as PPAR response elements, generally present in the promoter of genes. Binding to DNA and, thus, transcriptional activation is dependent upon formation of a heterodimer between PPAR and its indispensable partner retinoid X receptor, another member of the nuclear receptor superfamily.
The PPAR-isotype is mainly expressed in white adipose tissue (WAT) and regulates the expression of numerous genes involved in adipocyte differentiation and energy storage (2). PPARß/ is more widely expressed and has been connected with diverse functions ranging from regulation of fatty acid oxidation and inflammation to wound healing in skin (3).
PPAR is highly expressed in metabolically active tissues including liver, muscle, and brown adipose tissue. In hepatocytes, the liver cell type with the highest expression level of PPAR, PPAR governs lipid metabolism, gluconeogenesis, and amino acid metabolism (4).
In addition to governing metabolism, in recent years, it has also become evident that PPAR has an important role in regulating inflammatory responses in liver. By suppressing expression of proinflammatory genes PPAR controls and inhibits inflammation (5). One of the molecular mechanisms responsible for the immunosuppressive effects of PPAR is direct physical interaction with nuclear factor 
B (NF-B) (6), resulting in deactivation of this proinflammatory signaling pathway. Genes that are negatively regulated by PPAR in this fashion include acute phase genes and inflammatory signaling components like the IL-6 receptor (7).
Growing evidence has pointed to the involvement of a variety of inflammatory processes in the development of obesity and obesity-associated pathology (8). The recent demonstration that obesity is accompanied by a marked increase in macrophage infiltration of WAT has been instrumental in advancing our thoughts about the origin of inflammatory changes during obesity (9) (10). Indeed, it is now clear that obesity is strongly associated with an increase in circulating levels of acute phase proteins and cytokines, which mainly originate from WAT (11, 12). The influential role of inflammation in liver has been supported by studies showing that NF-B activation is a crucial step in the development of obesity-induced insulin resistance (13, 14).
Inasmuch as PPAR is able to suppress many of these inflammatory pathways, it might be an attractive target to reduce the inflammatory burden caused by obesity. However, so far, the role of PPAR and its ability to interfere with these inflammatory processes in liver have not been studied in the context of obesity-induced inflammation. To explore this function of PPAR in liver, wild-type (Wt) and PPAR–/– mice were chronically fed a low-fat diet (LFD) or high-fat diet (HFD). The HFD was given to induce moderate adiposity, resembling obesity. Our results indicate that the presence of PPAR in liver protects against chronic inflammation induced by chronically feeding a HFD.
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Materials and Methods |
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–/– mice and corresponding Wt mice were purchased at the Jackson Laboratory (Bar Harbor, ME). Male mice received a HFD or LFD for 6 months. The diets provided either 10 or 45% energy percent in the form of lard fat (D12450B or D12451; Research Diets, New Brunswick, NJ). Table 1
shows the composition of the diets. In another experiment, male Wt and PPAR–/– mice received Wy-14643 (Chemsyn Laboratories, Lenexa, KS) mixed in the food (0.1%) for 5 d. After both feeding experiments, liver and WAT were dissected, weighed, and directly frozen into liquid nitrogen. The animal experiments were approved by the animal experimentation committee of Wageningen University.
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Affymetrix GeneChip oligoarray analysis
Pooled RNA samples from five mice per experimental group were used for microarray analysis. Samples were hybridized on Affymetrix GeneChip Mouse Genome 430A arrays. Expression levels were calculated applying the multichip modified model for oligonucleotide signal (multi-mgMOS) (15) and a remapped Gene Chip Description (CDF) File (16). Heat maps were made in Spotfire DecisionSite software (Spotfire Inc., Sommerville, MA). Detailed descriptions of the applied methods are available on request.
Real-time PCR
RNA from animal tissue or cells was extracted with TRIzol reagent (Invitrogen) using the supplier’s instructions. One microgram of RNA was used for RT with the iScript cDNA Synthesis Kit (Bio-Rad Laboratories BV, Veenendaal, The Netherlands). Real-time PCR was done with platinum Taq polymerase (Invitrogen) and SYBR green using an iCycler PCR machine (Bio-Rad Laboratories BV). Melt curve analysis was included to assure a single PCR product was formed. The primers used are listed in Table 2.
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Liver triglycerides
Liver triglycerides were determined in 10% liver homogenates prepared in buffer containing 250 mM sucrose, 1 mM EDTA, 10 mM Tris-HCl (pH 7.5) using a commercially available kit from Instruchemie (Delfzijl, The Netherlands).
Immunohistochemistry
Cryosections of 5 µm from frozen liver were made. The coupes were dried overnight at room temperature followed by fixation in acetone for 5 min and acetone with 0.15% H2O2 for 5 min. For detection of macrophages/monocytes, an F4/80+ antibody (Serotec, Oxford, UK) was used. After preincubation with 20% normal goat serum, sections were incubated overnight at 4 C with the primary antibody diluted 1:50 in PBS/1% BSA. After incubation with the primary antibody, a goat antirat IgG conjugated to horseradish peroxidase (Serotec) was used as secondary antibody. Visualization of the complex was done using 3,3'-diaminobenzidene for 5 min. Negative controls were used by omitting the primary antibody. Oil-red O and hematoxylin and eosin staining of liver sections were done using standard protocols.
Primary hepatocyte isolation
Primary mouse hepatocytes from Wt and PPAR–/– mice were isolated as described previously (17). Briefly, after cannulation of the portal vein, the liver was perfused with calcium-free HBSS which was pregassed with 95% O2/5% CO2. Next, the liver was perfused with a collagenase solution (Sigma-Aldrich, Zwijndrecht, The Netherlands) until swelling and degradation of the internal liver structure was observed. The hepatocytes were released, filtered, and washed several times using Krebs buffer. The viability was assessed by using trypan blue (Sigma-Aldrich) and was around 80%. Cells were brought into culture using Williams E Medium supplemented with 10% FCS (Cambrex, Verviers, Belgium), penicillin/streptomycin/fungizone, insulin, and dexamethasone. Cells were plated on collagen (Serva Feinbiochemica, Heidelberg, Germany)-coated wells with a density of 0.5 x 106 cells/ml. After 4 h of incubation, the medium was removed and replaced with fresh medium. The next day, hepatocytes were used for experiments. IL-1ß and TNF were from R&D Systems Europe Ltd. (Abingdon, UK).
Isolation of adipocytes and stromal vascular cells
Freshly isolated epididymal adipose tissue was used for the isolation of adipocytes and stromal vascular cells. Minced adipose tissue was digested using collagenase (Sigma-Aldrich) at a concentration of 5 mg/ml dissolved in DMEM with 10% FCS. Tissues were incubated for 45 min at 37 C and were subsequently filtered through a 250-µM nylon mesh filter. After centrifugation, the floating cells were collected as adipocytes and the pelleted cells were collected as stromal vascular cells. Both cell fractions were washed with PBS, and RNA was isolated using TRIzol reagent (Invitrogen).
Immunoblot analysis
Immunoblotting was carried out using an ECL system (Amersham Biosciences, Diegem, Belgium) according to the manufacturer’s instructions. Equal amounts of liver cell lysates as determined by Bio-Rad Protein Assay reagent (Bio-Rad Laboratories BV) were resolved by SDS/PAGE on a 12% polyacrylamide gel. The F4/80+ antibody (Serotec) and the actin antibody (Sigma-Aldrich) were used at a dilution 1:1000 and the membranes were incubated overnight at 4 C. The secondary antibodies (goat antirat or rat antirabbit IgG, peroxidase; Sigma-Aldrich) were used at a dilution of 1:5000. All incubations were performed in 1x Tris-buffered saline, pH 7.5, with 0.1% Tween 20 and 5% dry milk. In the final washings, dry milk was removed from the solution.
Statistical analysis
Statistical significant differences were calculated using two-way ANOVA or Student’s t test. Correlations between gene expression signals and liver triglycerides content were assessed by Pearson’s correlation coefficient. The cutoff for statistical significance was set at a P value of 0.05 or below.
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Results |
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–/– vs. Wt mice fed a HFD). Changes in liver and adipose tissue weight in response to both diets were also evaluated. In comparison with LFD, HFD significantly increased adipose tissue to body weight ratio in Wt and PPAR–/– mice (Fig. 1B). In contrast, HFD increased liver to body weight ratio only in PPAR–/– mice (Fig. 1C). Overall, liver weights were higher in PPAR–/– mice.
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in obesity-induced hepatic inflammation, changes in gene expression in Wt and PPAR–/– mice after HFD were studied by Affymetrix GeneChip analysis. Wt and PPAR–/– mice showed increased expression of inflammatory genes in liver after HFD feeding compared with LFD feeding (Fig. 2). However, the effect of HFD feeding on inflammatory gene expression was more pronounced in the PPAR–/– animals compared with the Wt animals (Fig. 2). Genes that were highly increased in PPAR–/– mice fed the HFD included hepatic acute phase genes such as SAA and Orosomucoid, chemokines, and macrophage-related genes. These results suggest a higher degree of hepatic inflammation and the recruitment of proinflammatory cells including macrophages to the liver and indicate an antiinflammatory effect of PPAR in livers of mice fed the HFD.
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) and to study the expression of transcription factors and cytokines possibly involved in the regulation of these genes, qPCR analysis was performed. The majority of inflammatory genes that were analyzed showed an increased expression in the HFD group that was amplified in the PPAR–/– mice (Fig. 3). Expression of SAA, lipocalin, and CXCL10/IP-10 was 15-, 40-, and 25-fold higher, respectively, in PPAR–/– on HFD compared with Wt mice on LFD. Similar but somewhat mitigated changes were observed for metallothionein 2, vascular cell adhesion molecule-1 (VCAM-1), and several other genes. To assess which molecular pathways are implicated in the regulation of these genes, we analyzed the expression of several transcription factors and cytokines including STAT1 and 3, IL-6, IL-1ß, and TNF. Significant effects of both diet and genotype were only found for TNF. Together, these data point toward a protective effect of PPAR on obesity-induced hepatic inflammation.
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–/– mice fed the HFD–/– compared with the Wt mice, indicating more liver injury (Fig. 4A). In parallel with the gene expression results, in the HFD group, plasma SAA levels were markedly higher in the PPAR–/– compared with the Wt mice (Fig. 4B). The markedly elevated plasma levels of inflammatory marker SAA may point toward a chronic state of inflammation in the PPAR–/– animals after feeding the HFD (18).
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–/– vs. Wt mice fed the HFD–/– mice fed the HFD, suggesting an increase in inflammatory cell recruitment. qPCR analysis corroborated these data by showing markedly increased expression in PPAR–/– mice fed the HFD of two key genes implicated in macrophage/monocyte type cells recruitment: monocyte chemotactic protein 1 (MCP-1) and macrophage inflammatory protein 1 (MIP1) (Fig. 5). A similar expression pattern was observed for CD68 and F4/80+, two genes specifically expressed by macrophages. To determine changes in plasma concentrations of several chemokines, a multiplexing analysis was performed (Table 3): Except MIP1, all chemokines showed the highest plasma concentration in the PPAR–/– mice fed the HFD. In particular, plasma concentrations of MCP-1 and CXCL10/IP-10 were highly induced in HFD fed PPAR–/– mice. MCP-1 belongs to the subclass of CC chemokines, whereas CXCL10/IP-10 is part of the CXC subclass. Both subclasses are known to be involved in the recruitment of different inflammatory cell types (19, 20). Although the analysis of plasma concentrations of chemokines was performed on pooled plasma samples that precluded statistical analysis of the data, plasma concentrations of MCP-1 and CXL10/IP-10 perfectly fit with the hepatic gene expression data obtained by qPCR of individual mice.
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–/– mice fed the HFD was associated with macrophage infiltration, macrophages in liver were visualized using an antibody against F4/80+, a marker for mature macrophages (21). The number of activated macrophages present in the liver was found to be increased in the PPAR–/– mice compared with the Wt mice fed the HFD (Fig. 6A). No staining was observed in the absence of primary antibody (data not shown). Analysis of total liver cell lysates for F4/80+ protein content by immunoblot gave similar results (Fig. 6B). Together, these data indicate that the lack of PPAR promotes obesity-induced macrophage infiltration in liver.
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–/– mice on the LFD, which is consistent with the important function of PPAR in fatty acid catabolism (22). Remarkably, the combination of PPAR deletion and HFD feeding caused massive hepatic lipid accumulation, as shown by the appearance of large lipid droplets in histological sections of liver (Fig. 7A), which were visualized by Oil red O staining (Fig. 7B). Although Wt mice fed the HFD and PPAR–/– mice on LFD developed hepatic steatosis, PPAR–/– mice fed the HFD displayed clear signs of steatohepatitis with inflammatory cells including lymphocytes infiltrating the liver.
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–/– mice on HFD may be connected to hepatic lipid accumulation, we studied the correlation between inflammatory gene expression and hepatic lipid content. A highly significant positive correlation (P < 0.009–0.01, R = 0.83–0.56) was observed between the expression of numerous inflammatory genes and liver triglyceride content (Fig. 8) suggesting that the increase in triglyceride storage in the liver may be responsible for the elevated inflammatory status of the liver.
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is able to down-regulate expression of inflammatory genes in liver independent of its effect on hepatic lipid storage suggest that PPAR indirectly inhibits inflammation by preventing fat accumulation in liver. To examine whether PPAR might also directly suppress inflammatory gene expression independent of its influence on hepatic lipid storage, the effect of the PPAR agonist Wy-14643 on inflammatory gene expression in liver under nonsteatotic conditions was studied by Affymetrix GeneChip analysis. Mice were fed normal chow for 5 d with or without Wy-14643 mixed in their food. Whereas treatment of the mice with Wy-14643 did not result in significant changes in hepatic triglyceride content (Fig. 9A), numerous inflammatory genes were down-regulated by Wy-14643 in a PPAR-dependent manner (Fig. 9B). The results from microarray were confirmed for several genes by qPCR (Fig. 9C). Because not all genes analyzed after the HFD intervention are expressed under normal conditions, not all genes shown in Fig. 2 are presented here.
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on inflammatory gene expression in liver independent of its influence on hepatic lipid storage, the ability of PPAR to regulate genes involved in inflammatory cell recruitment was studied in primary mouse hepatocytes. Basal expression of MCP-1 and MIP1 was much higher in hepatocytes lacking PPAR, strengthening the possible involvement of PPAR in controlling the expression of these genes (Fig. 10). Furthermore, pharmacological PPAR activation using Wy-14643 resulted in decreased expression of MCP-1 and VCAM-1, changes which were not observed in hepatocytes cultured from PPAR–/–. To evaluate the role of PPAR in the regulation of these genes during inflammatory conditions, hepatocytes from both genotypes were treated with IL-1ß and TNF (Fig. 10). Expression of both cytokines was increased in liver of PPAR–/– mice after HFD feeding (see Fig. 3). Expression of MIP1 and VCAM-1 were much higher in PPAR–/– hepatocytes incubated in the presence of IL-1ß or TNF, respectively. In summary, these data suggest that PPAR is able to down-regulate the expression of numerous inflammatory genes in liver independent of its effect on hepatic lipid accumulation.
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governs inflammatory gene expression in adipose tissue–/– mice (Fig. 11A). The elevated expression of the macrophage marker gene F4/80+ in PPAR–/– mice on HFD was specific to the stromal vascular fraction of WAT (Fig. 11B), suggesting increased macrophage infiltration in the adipose tissue of the PPAR–/– mice after HFD feeding. The successful separation of stromal vascular cells from adipocytes is shown by the dominant expression of leptin in the adipocyte fraction. These data suggest that PPAR influences expression of inflammatory genes in adipose tissue, including cytokines, chemokines, and macrophage markers.
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Discussion |
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B is increased, which may lead to the development of insulin resistance (13).
The nuclear receptor PPAR has a major role in liver by altering the transcription of numerous target genes, many of which are involved in fatty acid oxidation. In line with its importance in fatty acid catabolism, positive effects of PPAR activation in the prevention and reversal of steatosis have already been demonstrated (25) (26). In addition to these diverse functions in hepatic metabolism, in the past decade a role for PPAR in controlling inflammation has clearly emerged (27, 28, 29). Since then, several molecular mechanisms by which PPAR exerts its antiinflammatory effects in liver and vascular wall have been uncovered (30, 31). Thus, although the connection between PPAR and inflammation is strong, little to no information is available yet on whether PPAR may modulate obesity-induced inflammation in liver. The aim of the present study was to determine whether PPAR may play a role in obesity-induced hepatic inflammation. Obesity was induced by chronically feeding Wt and PPAR–/– mice a HFD. Several lines of evidence suggest that PPAR protects against hepatic inflammation under conditions of obesity: 1) microarray and qPCR analysis indicated that expression of numerous genes involved in inflammation was markedly up-regulated in PPAR–/– mice fed a HFD compared with Wt mice fed a HFD. This included several acute phase genes and other inflammatory markers. 2) In plasma of PPAR–/– mice vs. Wt mice fed a HFD, markedly higher levels of SAA protein and ALT were measured, suggesting increased liver inflammation and injury. 3) Livers of PPAR–/– mice vs. Wt mice fed a HFD showed significantly increased infiltration of macrophages, as indicated by elevated presence of macrophage markers at the gene expression and protein level. These observations indicate that the presence of PPAR in liver protects against inflammation elicited by chronically feeding mice a HFD.
It can be theorized that PPAR may protect against obesity-induced hepatic inflammation by decreasing lipid storage in liver, which contributes to hepatic inflammation as observed in steatohepatitis. Alternatively, it is conceivable that PPAR suppresses the inflammatory response in liver by directly down-regulating the expression of target genes involved in inflammation. Finally, PPAR may act indirectly by suppressing inflammation in adipose tissue, thereby decreasing the secretion of adipokines that may promote hepatic inflammation. With respect to the former hypothesis, we observed markedly higher hepatic lipid storage in PPAR–/– vs. Wt mice fed a HFD, indicating the protective effect of PPAR against steatosis. When all mice were grouped together, highly significant correlations were observed between hepatic triglyceride concentration and gene expression levels of several acute phase proteins and cytokines, suggesting that the two are causally linked. These data would argue that PPAR influences obesity-induced hepatic inflammation mainly by decreasing lipid storage. However, we also observed that, under nonsteatotic conditions, either in vivo or in vitro, activation of PPAR by a synthetic agonist consistently down-regulated the expression of numerous inflammatory genes in a PPAR-dependent manner. This is in line with previous data showing a suppressive effect of PPAR on the expression of numerous genes implicated in hepatic inflammation in the absence of steatosis (32). Hence, PPAR suppresses inflammation regardless of changes in lipid storage in liver. Inasmuch as obesity is invariably connected with elevated hepatic lipid storage, and PPAR automatically decreases lipid storage by stimulating fatty acid catabolism, it is difficult to separate the relative contribution of these mechanisms to the overall effect of PPAR on obesity-induced hepatic inflammation.
Another possible mechanism directly implicates adipose tissue as the primary initiator of elevated hepatic inflammation. In mice fed the HFD, deletion of PPAR was associated with markedly elevated expression of numerous inflammatory genes in adipose tissue. This included cytokines, chemokines, and macrophage markers. Thus, it is possible that the elevated hepatic inflammation is secondary to events originating in the adipose tissue. These events are directly or indirectly governed by PPAR but are also independent of adiposity, as inflammatory gene expression was higher in HFD-fed PPAR–/– vs. Wt mice despite comparable fat mass. It should be emphasized that, although the expression of PPAR in adipose tissue is low compared with PPAR, this does not necessarily mean it is nonfunctional. Accordingly, it can be envisioned that, similar to its role in liver and the vascular wall, PPAR directly regulates expression of inflammatory genes in adipose tissue.
The development of chronic inflammation associated with obesity has partly been attributed to the infiltration of macrophages in WAT. Indeed, it was shown that, after 16 wk of HFD feeding, infiltration of macrophages occurred, resulting in elevated production of several proinflammatory mediators by WAT (9, 10). Whether similar obesity may also lead to increased macrophage infiltration in liver is not very clear. Our data suggest that the presence of PPAR prevents macrophage infiltration in liver in a mouse model of obesity. Although Kupffer cells represent the natural macrophage population in liver and play an important role in the immune defense, the higher expression levels of various chemokines and macrophage marker genes, including MCP-1, MIP1, CXCL10/IP-10, VCAM, and F4/80+, together with enhanced immunohistological staining for macrophage markers in the PPAR–/– mice fed the HFD, strongly suggests an increase in macrophage recruitment in liver (33, 34). As discussed above, PPAR may reduce macrophage infiltration by direct regulation of target genes involved in this pathway or indirectly by decreasing hepatic lipid storage, and by attenuating inflammation in adipose tissue.
We have previously shown that HFD feeding increases hepatic PPAR expression and activation, leading to the induction of classical PPAR target genes involved in fatty acid oxidation (35). The present data add a novel twist by showing that, whereas activation of PPAR down-regulates inflammatory gene expression, HFD increased inflammatory gene expression. The importance of PPAR is demonstrated by our finding that the increase in inflammatory gene expression caused by HFD becomes more dramatic in mice lacking PPAR. The observation that, in terms of regulation of inflammatory gene expression, the HFD-induced adiposity overrules the potential effect of HFD on PPAR activation attests to the notion that the effect of chronic HFD on inflammation is mediated by increased obesity.
In conclusion, PPAR exerts a marked antiinflammatory effect in liver in a mouse model of obesity, which appears to be at least partially achieved by decreasing activation and infiltration of inflammatory cells in liver. PPAR may reduce obesity-induced hepatic inflammation by diminishing fatty liver, which is tightly linked to elevated inflammatory status, by directly regulating inflammatory gene expression, or by suppressing inflammation in adipose tissue. Because PPAR automatically decreases steatosis via its effect on fatty acid catabolism, it is difficult to separate the relative contribution of these mechanisms to the overall effect of PPAR on obesity-induced hepatic inflammation.
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Acknowledgments |
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Footnotes |
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Abbreviations: ALT, Alanine aminotransferase; HFD, high-fat diet; LFD, low-fat diet; MCP-1, monocyte chemotactic protein 1; MIP1, macrophage inflammatory protein 1; NF-B, nuclear factor B; PPAR, peroxisome proliferator-activated receptor; qPCR, quantitative real-time PCR; SAA, serum amyloid A; VCAM-1, vascular cell adhesion molecule-1; WAT, white adipose tissue; Wt, wild type.
This study was supported by the Center for Human Nutrigenomics, with additional support by The Netherlands Organization for Scientific Research, the Dutch Diabetes Foundation, and the Wageningen Center for Food Sciences.
Disclosure Statement: The authors have nothing to disclose.
Received January 5, 2007.
Accepted for publication February 26, 2007.
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calls for activation in moderation: lessons from genetics and pharmacology. Endocr Rev 25:899–918 (PPAR) in the control of fatty acid catabolism. A new target for the treatment of metabolic syndrome. Biochimie (Paris) 86:833–837 target genes. Cell Mol Life Sci 61:393–416[CrossRef][Medline]B and PPAR. Adv Exp Med Biol 544:181–196[Medline] negatively regulates the vascular inflammatory gene response by negative cross-talk with transcription factors NF-B and AP-1. J Biol Chem 274:32048–32054 activator fenofibrate. J Biol Chem 279:16154–16160, leptin and IL-6 levels in obese women. Int J Obes Relat Metab Disord 28:993–997[CrossRef][Medline]B. Nat Med 11:183–190[CrossRef][Medline]-dependent hepatic lipid turnover in dietary steatohepatitis in mice. Hepatology 38:123–132[CrossRef][Medline] agonist, Wy-14,643, reverses nutritional fibrosis and steatohepatitis in mice. Hepatology 39:1286–1296[CrossRef]-leukotriene B4 pathway to inflammation control. Nature 384:39–43[CrossRef][Medline] protects against alcohol-induced liver damage. Hepatology 40:972–980[CrossRef][Medline]B and peroxisome proliferator-activated receptors in murine nonalcoholic fatty liver. Hepatology 40:376–385[CrossRef][Medline], ß/, and . J Clin Invest 114:1564–1576[CrossRef][Medline] activators in human C-reactive protein transgenic mice in vivo and in cultured human hepatocytes in vitro. Blood 103:4188–4194 mediates the effects of high-fat diet on hepatic gene expression. Endocrinology 147:1508–1516This article has been cited by other articles:
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