PPAR-δ agonist, GW501516 (GW), was obtained from Enzo Life Sciences (Farmingdale, NY, United States). Lipopolysaccharide (LPS) (Escherichia coli 0111:B4) and palmitic acid (PA) were purchased from Sigma-Aldrich (St Louis, MO, United States). Solutions and reagents used for cell culture were obtained from Invitrogen (Carlsbad, CA, United States) unless otherwise noted. Antibodies were purchased from Cell Signaling Technology Inc. (Beverly, MA, United States).

The animal protocol was designed to minimize pain or discomfort to the animals. Male 4-5-wk-old C57BL/6J mice were obtained from Japan SLC Inc. (Shizuoka, Japan). They were acclimatized to laboratory conditions (22-24 °C and 37%-64% humidity, with a 12-h dark-light cycle, free access to food and water) for one week prior to experimentation. Reflecting endotoxemia in NAFLD, we were decided to inject non-lethal very low dosage of LPS. Mice were randomly divided into four groups, which were treated for 12 wk as follows: standard diet (control, n = 5); HFD (HFD, 60% kcal from fat; D12492, Research Diets; New Brunswick, NJ, United States, n = 5); HFD plus one daily oral gavage of vehicle (0.5% carboxymethyl cellulose solution) with one weekly intraperitoneal (IP) injection of LPS (1 mg/kg per week) (HFD + LPS, n = 5); and HFD plus one daily oral dose of 3 mg/kg per day of GW501516, which was dissolved in the vehicle, with IP injection of LPS (HFD + LPS + GW, n = 6). Vehicle and GW501516 were administered for the last 3 wk. Body weight and food intake were recorded weekly. Body length (from nose to anus) was measured before sacrifice. Body mass index (BMI) was calculated by dividing body weight by the square of the body length (g/cm²)[22].

The study was reviewed and approved by the Institutional Review Board of Korea University and it was approved by the institutional animal review board of Korea University, Seoul, Korea, KUIACUC-2013-66 and conducted in compliance with the Guide for the Care and Use of Laboratory Animals.

The glucose tolerance test was conducted in all animals at 11 wk after dietary manipulation. After overnight fasting, 2 g/kg glucose was injected intraperitoneally, and blood samples were taken from the tail vein at 0, 15, 30, 60, 90, and 120 min. Blood glucose was measured using an Accu-Check Compact kit (Roche Diagnostics GmbH; Mannheim, Germany).

At 12 wk, the mice were intraperitoneally anesthetized with a mixture of tiletamine/zolazepam (30 mg/kg, Zoletil; Yuhan Corp.; Seoul, Korea) and xylazine (10 mg/kg, Rompun; Bayer, Inc.; Frankfurt, Germany), and sacrificed by exsanguination. Blood samples were extracted and the serum was isolated. The livers were rapidly excised and weighed. Serum alanine transaminase (ALT), aspartate transaminase (AST), triglyceride (TG), and total cholesterol (TC) levels were measured using common biochemical kits (Mindray Medical International Ltd.; Shenzhen, China).

The right liver lobe was stored at -80 °C until analysis of mRNA and protein. The left liver lobe was immediately fixed in 10% neutral-buffered formalin, paraffin-embedded, sectioned, and sections were stained with hematoxylin and eosin (HE). To visualize the neutral lipids, some of the frozen sections of fresh liver were stained using Oil Red O reagent. The liver samples were examined histologically in a blind manner by an experienced pathologist using the histological scoring system for NAFLD[23].

The human hepatoma HepG2 cell line (ATCC; Manassas, VA, United States) cells were cultured as manufacture’s instruction. In all experiments, PA concentration of 0.2 mmol/L which had no influence on cell viability was selected. The cells were stimulated with PA-BSA (0.2 μmol/L), LPS (1 μg/mL), or both, with or without GW501516 (1 or 10 μmol/L).

Total RNA was extracted from mouse liver tissues and HepG2 cells with TRIzol reagent according to the manufacturer’s instructions. RNA was reverse-transcribed into cDNA using the High Capacity cDNA Reverse Transcription kit (Applied Biosystems; Foster City, CA, United States). Gene expressions of NLRP1, NLRP3, NLRP6, NLRP10, NLRC4, caspase-1, and IL-1β were quantified by real-time polymerase chain reaction (PCR) with a 7000 Real-time PCR system (Applied Biosystems). PCR reactions were prepared with the Power SYBR Green PCR Master mix (Applied Biosystems) in 20 μL. The cycling parameters were 10 min at 95 °C followed by 40 cycles of 95 °C (15 s) and 30 s at 60 °C, followed by melting curve analysis. Gene expression by real-time PCR was presented relative to glyceraldehyde 3-phosphate dehydrogenase. The PCR primers used are listed in the Table 1.

Mouse liver tissues and HepG2 cell were lysed using M-PER Mammalian Protein Extraction Reagent (Pierce Chemical; Rockford, IL, United States). The protein concentration was determined using the bicinchoninic acid (BCA) method. Equal amounts (20 μg) of total proteins were boiled for 5 min and subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), followed by electrotransfer to polyvinylidene difluoride (PVDF) membranes. The membranes were blocked with 5% BSA in Tris-buffered saline (TBS) for 1 h at room temperature, followed by incubation with antibodies: anti-capase-1 (D7F10), anti-IL-1β (3A6), anti-p-AMPK-α (Thr172;40H9), anti-AMPK-α (23A3), and anti-β-actin (AC-15). The bands were visualized with an enhanced chemiluminescent direct labeling (ECL) system. The Gel Pro Analyzer 4.5, 2000 software (Media Cybernetics; Silver Spring, MD, United States) was used to determine the band density.

Liver was perfused with saline and homogenized in Tris-HCl buffer (20 mmol/L, pH 7.4). The homogenates were centrifuged at 2500 ×g for 10 min at 4 °C. Two hundred microliters of homogenates was analyzed for malondialdehyde (MDA) levels using a kit (Cell Biolabs, Inc.; San Diego, CA, United States), and the value was read at 532 nm.

Intracellular ROS levels in HepG2 cells were determined using 2′,7′-dichlorodihydrofluorescein diacetate (DCFDA). Cells were seeded on a 96-well bottom dishes. After the treatment described above in Cell culture and treatment, the cells were collected, and incubated with 10 μmol/L DCFDA (Molecular Probes; CA, United States) for 30 min at 37 °C. DCFDA fluorescence intensity was measured at 530 nm with an excitation wavelength of 488 nm using flow cytometry (Becton Dickinson; Heidelberg, Germany). The percentage of ROS-producing cells was determined by counting only those cells that produced high levels of ROS.

Data are presented as mean ± SD. Statistical significance was determined with one-way analysis of variance (ANOVA) followed by Tukey’s test for multiple comparisons using the GraphPad Instant program (GraphPad Software version 5.00; GraphPad Software Inc.; San Diego, CA). P < 0.05 was considered statistically significant. The statistical methods of this study were reviewed by a biomedical statistician.

The body weight and BMI of the HFD group were significantly higher compared with those of control mice (all P < 0.05) (Table 2). The food intake was greater in the HFD group compared with the HFD + LPS and HFD + LPS + GW501516 groups. The proportion of liver weight to body weight or body length was similar among the four groups. Serum AST and ALT levels were significantly increased in the HFD group compared with the control group (AST, 123.8 ± 30.54 IU/L vs 52.6 ± 10.33 IU/L; ALT, 136.6 ± 69.43 IU/L vs 23.4 ± 4.04 IU/L, all P < 0.05). GW treatment in HFD+LPS group significantly reduced serum AST and ALT compared with HFD (AST, 45.67 ± 11.11 IU/L vs 123.8 ± 30.54 IU/L; ALT, 20.5 ± 6.12 IU/L vs 136.6 ± 69.43 IU/L, all P < 0.05). HFD and/or LPS injection had no significant effect on TG or TC levels.

When subjected to a glucose tolerance test, glucose levels of control mice peaked at 30 min and returned to the basal level (Figure 1A). As expected, mice fed the HFD with or without LPS infection were more glucose-intolerant compared with controls, as demonstrated by the significant increase in the area under the curve (AUC) (53108 ± 9750 mg/dL and 52653 ± 9425 mg/dL vs 34545 ± 2776 mg/dL × 120 min, all P < 0.01) (Figure 1B). However, this glucose intolerance was not abolished by GW501516 treatment.

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Figure 1 Effects of high-fat diet for 12 wk on glucose tolerance in male mice treated with GW501516 (GW, 3 mg/kg per day for 3 wk). Intraperitoneal glucose tolerance test (A) and area under the curve for glucose (AUC_glucose) (B) showed glucose intolerance in mice fed the HFD with or without lipopolysaccharide (LPS) injection compared with the control group. GW treatment did not improve glucose intolerance. Data are expressed as the mean ± SD (5-6 mice per group). ^bP < 0.01 vs control group.

In addition to steatosis, chronic inflammation is an important contributing factor in NASH pathogenesis[5]. To investigate the inflammatory response in this model, we measured hepatic caspase-1 and IL-1β levels. The gene expression level of caspase-1, which is known to be activated by the NLRP3 inflammasome complex and to cleave pro-IL-1β to the active form, was increased in the HFD + LPS group relative to the control group (P < 0.01) (Figure 2A). The hepatic gene expression and protein levels of IL-1β were also increased in the HFD + LPS group relative to the control group (P < 0.05 for mRNA expression and P < 0.001 for protein-level expression) (Figure 2B and C). When compared with HFD+LPS, PPAR-delta agonist GW treatment reduced hepatic caspase-1 mRNA (2.8-fold vs 1.7-fold) and IL-1β mRNA (2.2-fold vs 1.6-fold).

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Figure 2 Effects of GW501516 on caspase-1 and IL-1β levels in mice fed a high fat diet. Hepatic mRNA levels of caspase-1 (A) and IL-1β (B) at 12 wk showed a significant increase in the HFD + lipopolysaccharide (LPS) group compared with the control group. GW501516 treatment tended to ameliorate this increase. Representative immunoblot and graphic presentation show hepatic protein levels of IL-1β (C) in different groups of mice as indicated. Mice fed an HFD with LPS injection with or without GW501516 treatment had increased protein levels of IL-1β. Data are expressed as the mean ± SD (5-6 mice per group). ^aP < 0.05, ^bP < 0.01, and ^dP < 0.001 vs control group.

We also investigated the effect of PPAR-δ activator on inflammasome induced by PA and LPS in HepG2 cells. First of all, we analyzed the PPAR-δ protein expression itself in various stimuli. In vitro exposure with- or without stimuli such as LPS only, PA only or GW only, or combined with it, PPAR-delta protein expressions were not different. After then, we analyzed the expression of several inflammasome-related mRNAs upon activation with PA and/or LPS. PA and LPS together elicited the mRNA expression of NLRP3, NRLP6, and NLRP10 (all P < 0.05) (Figure 3). We next tested whether GW501516 could prevent the overexpression of such inflammasome components and the downstream pro-inflammatory signals. As shown in Figure 4, treatment with 10 μmol/L GW501516 reduced the PA and LPS-induced mRNA expression of NLRP3, NRLP6, and NLRP10 (all P < 0.05). GW501516 also reduced the expression of caspase-1 and IL-1β mRNA in a dose-dependent manner (all P < 0.05) (Figure 4D and E). In concordance with mRNA expression, the presence of GW501516 reduced the PA and LPS-induced production of caspase-1 and IL-1β protein expression (Figure 4F).

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Figure 3 Expression of inflammasome components in HepG2 cells after stimulation with palmitic acid and lipopolysaccharide. Relative mRNA levels of NLRP1, NLRP3, NLRC4, NLRP6, and NLRP10 in HepG2 cells were analyzed by RT-PCR. Compared with the control group, significantly increased expression of NLPR3, NLRP6, and NLRP10 was observed in the PA and LPS stimulation group. Data are expressed as the mean ± SD. ^aP < 0.05 vs control cells.

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Figure 4 Effects of GW501516 on palmitic acid- and lipopolysaccharide-induced inflammasome and pro-inflammatory cytokine in HepG2 cells. Relative mRNA levels of NLRP3 (A), NLRP6 (B), NLRP10 (C), caspase-1 (D), and IL-1β (E) in HepG2 cells were analyzed by RT-PCR. GW501516 inhibited PA- and LPS-induced mRNA expression of several inflammasome components, caspase-1, and IL-1β. Representative immunoblot and graphic presentation show protein levels of caspase-1 and IL-1β (F) in HepG2 cells. PA and LPS tended to elicit caspase-1 and IL-1β release, and GW501516 reduced this processing significantly. Data are expressed as the mean ± SD. ^aP < 0.05 vs control cells, ^cP < 0.05 and ^bP < 0.01 vs PA + LPS-treated cells.

Histological images of liver pathology were obtained. Liver sections were stained with HE and Oil Red O. In mice fed the standard diet, there was no detectable fatty change in the microscopic image. By contrast, relative to the control group, mice on a 12-wk HFD with or without LPS injection developed extensive macrovesicular steatosis (0.0 vs 17.0% ± 5.7% and 16.4% ± 14.7%, all P < 0.05) and inflammation around the perisinusoidal area. Ballooned hepatocytes and fibrosis were not observed in any study group. In mice that were administrated GW501516, the intensity of fat accumulation (2.7% ± 2.6%) and NAS (0.5) were significantly decreased (P < 0.05) compared to those of the HFD + LPS group (16.4% ± 14.7% and 1.8%, respectively) (Figure 5A, B and C).

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Figure 5 Histopathological features of livers in mice fed a high fat diet with or without lipopolysaccharide injection and GW501516 treatment. A: Hematoxylin and eosin (HE) staining and Oil Red O staining of hepatic lipid accumulation (magnification × 100). In mice fed an HFD, moderate macrovesicular steatosis (17.0%) and inflammatory cell infiltration were observed compared with the control group. The macrovesicular steatosis was improved to 2.7% following GW501516 treatment; B: Histogram of the percentage of hepatocytes showing macrovesicular fatty change; C: Nonalcoholic fatty liver disease (NAFLD) activity score. The NAFLD activity score in the GW501516-treated group was significantly lower than that in the HFD + LPS group (1.8 vs 0.5, P < 0.05). Data are expressed as the mean ± standard deviation (SD) (5-6 mice per group). ^aP < 0.05, ^bP < 0.01, and ^dP < 0.001 vs control group. ^cP < 0.05 vs the HFD + LPS group.

Oxidative stress is thought to play a role in the pathogenesis of NASH, and ROS are essential for inflammasome activation[7,14]. A large body of evidence indicates that AMP-activated protein kinase (AMPK) is an essential regulator of fatty acid metabolism[21], and suppresses ROS generation by regulating intracellular nicotinamide adenine dinucleotide phosphate (NADPH) production[18,24]. In the in vivo study, levels of MDA, a product of lipid peroxidation, in liver homogenates, there was no statistically significant difference in four groups (Figure 6A). We also examined the effect of GW501516 on AMPK activation. As the activity of AMPK correlates with phosphorylation at Thr-172, the activation of AMPK was assessed by determining phosphorylation of AMPKα. As shown in Figure 6B, treatment with GW501516 increased the levels of AMPKα phosphorylation in mice fed the HFD with LPS injection by 1.6-fold compared to the control group.

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Figure 6 Effects of GW501516 on lipid peroxidation and associated molecular pattern-activated protein kinase-α phosphorylation in mice fed a high fat diet. Liver malondialdehyde (MDA) levels (A) showed a slight increase in the HFD + lipopolysaccharide (LPS) group without statistical significance; B: Representative immunoblot and graphic presentation of p-AMPK-α and total AMPK-α protein levels in the livers of mice are shown. Treatment of GW501516 enhanced phosphorylation of AMPK-α in mice fed an HFD with LPS injection. Data are expressed as the mean ± SD (5-6 mice per group).

In the in vitro study, the effect of GW501516 on PA and LPS-induced ROS production was investigated using DCFDA to detect cellular ROS levels. As illustrated in Figure 7A, incubating HepG2 cells with LPS and LPS + PA induced a significant increase in the cellular ROS level at 16 h (all P < 0.05 vs control group). Treatment with GW501516 (1 μmol/L and 10 μmol/L) attenuated the ROS generation in the PA- and LPS-stimulated HepG2 cells (all P < 0.05 vs PA + LPS group). In addition, we assessed AMPK activation with PA and LPS. PA or LPS alone had little effect on phosphorylation of the AMPKα subunit. However, adding GW501516 (1 μmol/L and 10 μmol/L) substantially increased phosphorylation of the AMPKα subunit in HepG2 cells (2.3- and 2.2-fold vs control group) (Figure 7B). These in vivo and in vitro results demonstrated that GW501516 could suppress hepatic oxidative stress and enhance phosphorylation of the AMPKα under high fat conditions.

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Figure 7 Effects of GW501516 on reactive oxygen species production and associated molecular pattern-activated protein kinase-α phosphorylation in HepG2 cells. Intracellular ROS production was quantified using the fluorescent probe 2′,7′-dichlorodihydrofluorescein diacetate (DCFDA) (A). GW501516 inhibited lipopolysaccharide (LPS)- and palmitic acid (PA)-induced ROS generation. Representative immunoblot and graphic presentation of p-AMPK-α and total AMPK-α protein levels in HepG2 cells are shown (B). GW501516 tended to increase the phosphorylation of AMPK-α in PA- and LPS-treated cells. Data are expressed as the mean ± SD (5-6 mice per group). ^aP < 0.05 vs control cells, ^cP < 0.05 vs PA + LPS-treated cells.

Nonalcoholic fatty liver disease (NAFLD) is one of the most common disease of the liver. Although thiazolidinediones and antioxidants such as vitamin E have been evaluated in several clinical trials, few pharmacological treatments can be recommended at present. Inflammasome is a large, intracellular multi-protein complex that is a sensor of pathogen-associated molecular patterns (PAMPs) or damage-associated molecular patterns (DAMPs) that govern the cleavage of pro-inflammatory cytokines such as pro-interleukin (IL)-1β and pro-IL-18. Previous studies have shown that NLRP3 inflammasome is involved in the pathogenesis of obesity-induced inflammation, insulin resistance, and development of type 2 diabetes. However, the role of inflammasome in the pathogenesis of NAFLD/NASH has not been elucidated.

Peroxisome proliferator-activated receptors (PPARs) are members of the nuclear receptor superfamily, and three isotypes exist in mammals. It has become increasingly evident that PPAR-δ is also an important metabolic regulator, as its activation enhances fatty acid oxidation, improves glucose homeostasis, and attenuates macrophage inflammatory responses. Current research hot spot is to whether the treatment with the PPAR-δ agonist could ameliorate high fat-induced inflammasome activation and the inflammatory response using in vivo and in vitro NAFLD models.

Until now, studies on the relationship between PPAR-δ activation and inflammasome have been scarce. In a recent study, the PPAR-δ agonist GW0742 was shown to attenuate the renal dysfunction and inflammation caused by chronic high-fructose corn syrup-55 exposure by preventing activation of the NLRP3 inflammasome in the kidney. To investigate whether the treatment with the PPAR-δ agonist could ameliorate high fat-induced inflammasome activation and the inflammatory response in the liver, the authors were using in vivo and in vitro NAFLD models. This study showed that PPAR-δ agonist GW501516 was found to reduce the activation of inflammasome as well as the overproduction of pro-inflammatory cytokines in HepG2 cells and the livers of mice. Moreover, GW501516 alleviated hepatic steatosis in vivo.

The study results suggested that the PPAR-δ agonist GW501516 suppresses the activation of inflammasome and reduces IL-1β levels in the liver, possibly by modulation of AMPK phosphorylation and decreased production of ROS. This anti-inflammatory effect might be associated with improvement of hepatic steatosis in mice. The targeting of inflammasome by the PPAR-δ agonist may have therapeutic implications for the treatment of NAFLD.

NAFLD is a spectrum of disease entity including simple steatosis, steatosis with inflammation, fibrosis and cirrhosis. Inflammasome is a large, intracellular multi-protein complex that is a sensor of PAMPs or DAMPs that govern the cleavage of pro-inflammatory cytokines such as pro-IL-1β and pro-IL-18.

The authors studied effects of PPAR-δ activator; GW501516 on inflamasome pathway in mouse model of NASH (high fat diet, LPS/PA) and HepG2 cells culture. They show that the beneficial role in mouse liver was through increased NLRP3-10 in HepG2 cells.