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British Journal of Pharmacology logoLink to British Journal of Pharmacology
. 2017 Jun 9;174(14):2358–2372. doi: 10.1111/bph.13844

Hydroxyeicosapentaenoic acids and epoxyeicosatetraenoic acids attenuate early occurrence of nonalcoholic fatty liver disease

Chunjiong Wang 1,†, Wenli Liu 1,†, Liu Yao 1,†, Xuejiao Zhang 1, Xu Zhang 1, Chenji Ye 1, Hongfeng Jiang 2, Jinlong He 1, Yi Zhu 1, Ding Ai 1,✉
PMCID: PMC5481642  PMID: 28471490

Abstract

Background and Purpose

The ω‐3 polyunsaturated fatty acids (PUFAs) mediate protective effects on several metabolic disorders. However, the functions of their metabolites in the early stage of nonalcoholic fatty liver disease (NAFLD) are largely unknown.

Experimental Approach

Mice were fed a control diet, high‐fat diet (HFD) or ω‐3 PUFA‐enriched HFD (ω3HFD) for 4 days and phenotypes were analysed. LC–MS/MS was used to determine the eicosanoid profiles. Primary hepatocytes and peritoneal macrophages were used for the mechanism study.

Key Results

In short‐term HFD‐fed mice, the significantly increased lipid accumulation in the liver was reversed by ω‐3 PUFA supplementation. Metabolomics showed that the plasma concentrations of hydroxyeicosapentaenoic acids (HEPEs) and epoxyeicosatetraenoic acids (EEQs) were reduced by a short‐term HFD and markedly increased by the ω3HFD. However, HEPE/EEQ treatment had no direct protective effect on hepatocytes. ω3HFD also significantly attenuated HFD‐induced adipose tissue inflammation. Furthermore, the expression of pro‐inflammatory cytokines and activation of the JNK pathway induced by palmitate were suppressed by HEPEs and EEQs in macrophages. 17,18‐EEQ, 5‐HEPE and 9‐HEPE were identified as the effective components among these metabolites, as indicated by their greater suppression of the palmitate‐induced expression of inflammatory factors, chemotaxis and JNK activation compared to other metabolites in macrophages. A mixture of 17,18‐EEQ, 5‐HEPE and 9‐HEPE significantly ameliorated the short‐term HFD‐induced accumulation of macrophages in adipose tissue and hepatic steatosis.

Conclusion and Implications

17,18‐EEQ, 5‐HEPE and 9‐HEPE may be potential approaches to prevent NAFLD in the early stage by inhibiting the inflammatory response in adipose tissue macrophages via JNK signalling.

Abbreviations

ACC

acetyl‐CoA carboxylase

ARA

arachidonic acid

CCL

C‐C motif chemokine ligand

DHA

docosahexaenoic acid

EEQ

epoxyeicosatetraenoic acid

EPA

eicosapentaenoic acid

FAS

fatty acid synthase

FFA

free fatty acid

HEPE

hydroxyeicosapentaenoic acid

HFD

high‐fat diet

PUFA

polyunsaturated fatty acid

TG

triglyceride

ω3HFD

ω‐3 PUFA‐enriched HFD

Introduction

Nonalcoholic fatty liver disease (NAFLD) is a major health burden worldwide. Obesity is an independent risk factor for a number of metabolic disorders, including NAFLD (Li et al., 2016). Environmental factors, especially a high consumption of fat, carbohydrates and protein, are the main reasons for the high prevalence of obesity (Siddiquee et al., 2015). Within 1 week, a high‐fat diet (HFD) can induce adipose inflammation, NAFLD and insulin resistance in both rodents and humans (Brons et al., 2009; Wiedemann et al., 2013; Senthil Kumar et al., 2014; Boon et al., 2015; Wulan et al., 2015). Therefore, the early stage of high‐fat consumption is a crucial part of the pathological process of metabolic disorders, but this has received less attention than chronic obesity.

The ω‐3 polyunsaturated fatty acids (PUFAs), including eicosapentaenoic acid ( EPA) and docosahexaenoic acid ( DHA), have protective roles in NAFLD. ω‐3 PUFAs can decrease the alanine aminotransferase level, serum lipid levels and hepatic lipid accumulation in patients with NAFLD (Capanni et al., 2006; Zhu et al., 2008). Similarly, in rodents, ω‐3 PUFA supplementation and endogenously synthesized ω‐3 PUFA in Caenorhabditis elegans fat‐1 transgenic mice ameliorated long‐term HFD‐induced fatty liver, insulin resistance and hyperlipidaemia (Kim et al., 2012; Espinosa et al., 2013; Li et al., 2014; Soni et al., 2015). However, the effect of ω‐3 PUFA on the onset of NAFLD and the underlying mechanism are still unknown.

EPA and DHA are present in phospholipids of the cell membrane and share the same enzymes with arachidonic acid (ARA) in the eicosanoid‐producing process. EPA and DHA are metabolized into numerous eicosanoids by COX, lipoxygenase and cytochrome P450. Among these metabolites, resolving E1 significantly ameliorated fatty liver disease and insulin resistance in db/db mice, and protectin D1 up‐regulated the expression of adiponectin in adipose tissue (Gonzalez‐Periz et al., 2009). These results indicate the putative protective roles of eicosanoids derived from ω‐3 PUFA in NAFLD. However, the metabolites of ω‐3 PUFA are numerous and complex, and their functions in NAFLD are largely unknown.

Because ω‐3 PUFAs can compete the same metabolic enzymes with the ω‐6 type, ω‐3 PUFA supplementation may also affect the metabolic profiles of ARA, thereby shifting ARA eicosanoid metabolism. Thus, determining the metabolic profile of both ω‐3 and ω‐6 PUFAs may be important to understand the protective role of ω‐3 PUFA in NAFLD.

In our previous study, we developed an LC–MS/MS‐based metabolomic method analysing 32 ARA metabolites and 37 ω‐3 PUFA‐derived products (Zhang et al., 2015). In the present study, using this method, we have determined the changes in metabolic profiles of ω‐3 and ω‐6 PUFAs in the plasma of mice after short‐term (4 day) HFD treatment, with or without ω‐3 PUFA supplementation, to screen the putative metabolites preventing the onset of NAFLD. We also studied the mechanisms. ω‐3 PUFA supplementation ameliorated the short‐term HFD‐induced inflammatory response in adipose tissue and, consequently, hepatic steatosis. By using targeted metabolomics, we demonstrated that 17,18‐epoxyeicosatetraenoic acid (EEQ), 5‐hydroxyeicosapentaenoic acid (HEPE) and 9‐HEPE derived from EPA mediated the protective effects of ω‐3 PUFA.

Methods

Experimental animals and diets

Specific‐pathogen‐free C57BL/6 mice were obtained from the Experimental Animal Centre of Military Medical Science Academy, Beijing, China. Eight‐week‐old male C57BL/6 mice (21–24 g) were randomized into groups. The investigator was not blind to the experimental groups. To study the effect of ω‐3 PUFA on short‐term HFD‐induced metabolic disorders, mice were divided into three groups for treatment: control diet (normal chow, 10% fat), HFD (45% fat) or ω‐3 PUFA‐enriched HFD (ω3HFD) (45% fat and 3% wt.wt‐1 ω‐3 PUFA) for 4 days (Wu et al., 2013). To study the effect of ω‐3 PUFA on the normal condition, mice were treated with the control diet or ω‐3 PUFA‐enriched diet (3% wt.wt‐1 ω‐3 PUFA in control diet). To study the in vivo functions of ω‐3 PUFA metabolites, control‐diet and HFD‐treated mice were injected i.p. with a mixture of 17,18‐EEQ, 5‐HEPE and 9‐HEPE with equal amounts of each every other 12 h for 4 days (70 ng·g−1 body wt). I.p. glucose tolerance testing (IPGTT) was performed on day 3, and mice were killed by exsanguination after being anaesthetized with isoflurane on day 4. The food intake was monitored throughout the feeding period. All mice were housed in a temperature‐controlled environment in individually ventilated cages with wood shavings as bedding (3–6 per cage) with 12 h light/dark cycles and received food and water ad libitum. All procedures involving experimental animals were performed under the principle for replacement, refinement or reduction (the 3Rs) and in accordance with the US National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Publication No. 85‐23, updated 2011) and were approved by the Institutional Animal Care and Use Committee of Tianjin Medical University (Tianjin, China). Animal studies are reported in compliance with the ARRIVE guidelines (Kilkenny et al., 2010; McGrath and Lilley, 2015).

Sample preparation for LC–MS/MS

Plasma and hepatic tissue were prepared for LC–MS/MS as described previously (Zhang et al., 2015). In brief, plasma was extracted by solid‐phase extraction (SPE). Before extraction, Waters Oasis‐HLB cartridges were conditioned with methanol and Milli‐Q water. Samples (200 μL) containing internal standard (IS) mixture (5 ng for each) were loaded onto cartridges, which were washed with 1 mL 5% methanol. The aqueous plug was pulled from the SPE cartridges under high vacuum, and SPE cartridges were further dried under high vacuum for 20 min. Analytes were eluted into tubes with 1 mL methanol. The eluent was then evaporated to dryness under a nitrogen stream.

Mouse livers were extracted by liquid–liquid extraction. After being weighed, tissues were homogenized with 500 μL methanol (2% formic acid and 0.01 mol·L−1 butylated hydroxytoluene) spiked with IS mixture. Samples were vigorously mixed on a vortexer for 5 min. After centrifugation (12 000× g for 10 min at 4°C), the supernatant was transferred to a new tube. Water (700 μL) and ethyl acetate (1 mL) were added. The sample was mixed vigorously for 2 min and centrifuged for 10 min at 12 000× g. The upper organic phase was transferred to a new tube, and the water phase was extracted again. The organic phase was combined and then evaporated to dryness.

The dried residue extracted by SPE and liquid–liquid extraction was dissolved in 100 μL 30% acetonitrile. After vigorous mixing, samples were filtered by use of centrifuge tube filters (nylon membrane, 0.22 μm).

Ultraperformance liquid chromatography

Chromatographic separations involved use of an ultraperformance LC (UPLC) BEH C18 column (1.7 μm, 50 × 2.1 mm i.d.) consisting of ethylene‐bridged hybrid particles. The column was maintained at 25°C, and the injection volume was set to 10 μL. The mobile‐phase flow rate was 0.6 mL·min−1. The gradient was 0–1.5 min from 30 to 40% neat acetonitrile; 1.5–6.5 min to 60% neat acetonitrile; 6.5–7.6 min to 80% neat acetonitrile, which was maintained for 1 min; and 8.6–8.8 min reduced to 30% neat acetonitrile and maintained for 0.2 min. The same gradient was applied to 37 ω‐3 PUFA metabolites (Zhang et al., 2015).

Mass spectrometry

Targeted profiling of ω‐6 and ω‐3 PUFA metabolites involved the use of a 5500 QTRAP spectrometer equipped with a turbo ion spray electrospray ionization source. Analytes were detected by multiple reaction monitoring (MRM) scans in negative mode. The MRM transitions for each eicosanoid were as described previously (Zhang et al., 2015). The dwell time used for all MRM experiments was 25 ms. The ion source parameters were CUR = 40 psi, GS1 = 30 psi, GS2 = 30 psi, IS = −4500 V, CAD =medium and temperature = 500°C (Zhang et al., 2015). The experiments and data analysis of LC–MS/MS were carried out in a blind manner. In Figure 2F, the level of metabolites was normalized to that of control group to show the fold change of every metabolite clearly, and the concentrations of these metabolites are shown in Table S2.

Oil red O staining

Liver samples were fixed with 4% paraformaldehyde and embedded in Optimal Cutting Temperature compound for frozen sectioning. Liver sections were stained with Oil red O staining solution for 30 min. After being rinsed with PBS, nuclei were stained with haematoxylin.

Determination of hepatic triglyceride and total cholesterol content

In total, 50 mg liver tissue was homogenized at 4°C in 1 mL chloroform‐methanol extraction buffer (2:1). After extraction for 16 h in 4°C, samples were neutralized with 300 μL deionized water and then centrifuged at 13 000 × g for 10 min. The subnatant was collected and blow‐dried by nitrogen. After the samples had been thawed with 200 μL 5% TritonX‐100, triglyceride (TG) or total cholesterol content was determined by using a TG or cholesterol determination kit according to the manufacturer's instructions.

Determination of the concentration of inflammatory compounds in plasma

The concentrations of inflammatory agents in plasma were determined by the BD cytometric bead array mouse inflammation kit (BD, catalogue no. 552364).

Culture of primary mouse hepatocytes

Male C57BL/6 mice (16–18 g) 5 to 6 weeks old were anaesthetized with isoflurane; then a catheter was placed in the inferior vena cava. The liver was perfused with heparin, 40 mL solution I (Krebs's solution +0.1 mM EGTA) and 30 mL solution II (Krebs's solution +2.74 mM CaCl2 + 0.05% collagenase I). The perfused liver was passed through a 400 screening‐size filter (38 μm diameter) by flushing with RPMI 1640 medium. The hepatocytes were collected by centrifugation at 50× g for 2 min, re‐suspended with RPMI 1640 medium and placed in six‐well plates for experiments after three washes with RPMI 1640.

Macrophage culture

Peritoneal macrophages were harvested by peritoneal lavage 3 days after an i.p. injection of 3% thioglycollate and cultured at 37°C in DMEM containing 10% FBS (Jiang et al., 2014).

Chemotaxis assay of macrophages

Mice were fed a chow diet, HFD or ω3HFD for 4 days. Then 50 mg adipose tissue was removed and cultured in the lower Transwell chamber containing a polycarbonate filter with pore size 8 μm in DMEM for 12 h. Macrophages were seeded in the upper chamber. After 4 h, cells were fixed in formalin and stained with DAPI. Migrating macrophages on the bottom of filters were counted.

For the in vitro chemotaxis assay, macrophages were cultured with palmitate and the indicated metabolites for 24 h in the lower Transwell chamber. Untreated macrophages were seeded in the upper chamber. After 4 h, cells were fixed in formalin and stained with DAPI. Migrating macrophages on the bottom of filters were counted.

qPCR assay

Total RNA extracted from cells and tissues was reverse‐transcribed to cDNA for qPCR. Each sample was measured in duplicate or triplicate. To control for unwanted sources of variation, we normalized the relative expression of the target genes in various groups to that of β‐actin and calculated the expression by the 2−ΔCt methodology (Wang et al., 2014). qPCR followed the Minimum Information for Publication of Quantitative Real‐Time PCR Experiments guidelines. The sequences of primers for real‐time PCR are shown in Table S1.

Western blot analysis

Total protein was extracted and separated by 10% or 12% SDS‐PAGE and then transferred to a PVDF membrane, which was immunostained with primary antibodies for p‐JNK, JNK, p‐p38 MAPK (p38), p38, p‐ERK or ERK at 4°C overnight. To control for unwanted sources of variation, we normalized the relative expression of target proteins in various groups to that of β‐actin.

Statistical analysis

Data are presented as mean ± SEM, and two or more groups were compared by Student's t‐test or one‐way ANOVA by using GraphPad Prism 5 (GraphPad software). For one‐way ANOVA, Bonferroni's multiple comparison post hoc test was performed for data with F at P < 0.05 and no significant variance inhomogeneity. Pearson correlation was used to assess correlation by using R v3.0.3 (packages muma and GMD) (Zhao and Sandelin, 2012). Statistical significance was set at P < 0.05. The data and statistical analysis comply with the recommendations on experimental design and analysis in pharmacology (Curtis et al., 2015).

Materials

Control diet, ω‐3 PUFA‐enriched diet, HFD and ω3HFD were from Medicience (Yangzhou, China). HEPEs and EEQs were from Cayman Chemical (Ann Arbor, MI, USA). UPLC BEH C18 column consisting of ethylene‐bridged hybrid particles was from Waters (Milford, MA, USA). 5500 QTRAP spectrometer was from AB Sciex (Foster City, CA, USA). Transwell chamber was from Corning (NY, USA). TG and total cholesterol determination kits were from BioSino Bio‐Technology and Science (Beijing). BD cytometric bead array mouse inflammation kit was from BD Biosciences (Franklin Lakes, New Jersey, USA). Antibodies for p‐JNK, JNK, p‐p38, p38, p‐ERK or ERK were from Cell Signaling Technology (Danvers, MA, USA). Antibody for MAC‐3 was from Santa Cruz Biotechnology (Dallas, TX, USA).

Nomenclature of targets and ligands

Key protein targets and the ligands in this article are hyperlinked to corresponding entries in http://www.guidetopharmacology.org, the common portal for data from the IUPHAR/BPS Guide to PHARMACOLOGY (Southan et al., 2016), and are permanently archived in the Concise Guide to PHARMACOLOGY 2015/2016 (Alexander et al., 2015a,b,c).

Results

ω‐3 PUFA ameliorated short‐term HFD‐induced hepatic steatosis and dyslipidemia

To determine the effect of ω‐3 PUFA on short‐term HFD‐induced hepatic steatosis, we added ω‐3 PUFA in the HFD (ω‐3HFD). Mice were fed the control diet, HFD or ω3HFD for 4 days. The daily food intake was decreased in mice with HFD feeding compared with the control diet, but ω‐3 supplementation had no additive effect as compared with the HFD alone (Figure S1A). IPGTT revealed that ω‐3 PUFA had no effect on short‐term HFD‐induced glucose intolerance (Figure S1B). The body, liver, fat and heart weights, and the ratios of liver to body weight, fat to body weight and heart to body weight were comparable among the three groups (Figure S1C–E). The morphology of livers from mice subjected to short‐term HFD or ω3HFD did not differ on haematoxylin and eosin staining (Figure 1A). On oil red O staining, the hepatic steatosis induced by the HFD was greatly ameliorated by ω‐3 PUFA (Figure 1A), which agreed with a significant reduction in hepatic TG content (Figure 1B). Hepatic total cholesterol content was not changed by the HFD (Figure 1C). In contrast, the HFD significantly increased plasma total cholesterol and LDL‐cholesterol level, which was suppressed by ω3HFD (Figure 1D, E). However, ω‐3 PUFA supplementation in the control diet had no effect on hepatic TG or total cholesterol content or plasma cholesterol levels (Figure S2A–E).

Figure 1.

Figure 1

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ω‐3 PUFA ameliorated short‐term HFD‐induced hepatic steatosis and dyslipidaemia. Mice were fed a control diet (Ctrl), HFD or ω3HFD for 4 days. (A) Oil red O staining (ORO) (upper panel) of hepatic lipid accumulation and haematoxylin and eosin (H&E) staining (lower panel) of liver morphology. (B, C) Quantitative assay of TGs (B) and total cholesterol (CHO) (C) content in livers. (D, E) Plasma CHO (D) and LDL‐CHO (LDL‐C) (E) levels in Ctrl, HFD and ω3HFD mice. (F) qPCR analysis of mRNA levels of genes involved in lipid metabolism. The mean values of the Ctrl group were set to 1. The values of other groups were normalized to Ctrl group values, represented as fold of Ctrl values. n = 13 animals per group. Data are mean ± SEM. *P < 0.05 versus Ctrl, #P < 0.05 versus HFD. SREBP1, sterol regulatory element binding protein‐1; ChREBP, carbohydrate‐responsive element‐binding protein; SCAD, short‐chain fatty acyl CoA dehydrogenase; MCAD, medium‐chain fatty acyl CoA dehydrogenase; AOX1, acyl‐CoA oxidase; LDLR, low density lipoprotein receptor.

We measured the expression of genes involved in hepatic lipogenesis, fatty acid oxidation, lipid uptake and lipoprotein secretion. As compared with the HFD, ω3HFD significantly decreased the expression of acetyl‐CoA carboxylase (ACC) and fatty acid synthase (FAS) and increased that of acyl‐CoA oxidase and short‐chain and medium‐chain fatty acyl‐CoA dehydrogenase (Figure 1F). These findings suggest that decreased lipogenesis and increased β oxidation may be involved in the protective role of ω‐3 PUFA in the early phase of HFD‐induced hepatic steatosis.

ω‐3 PUFA changed eicosanoid profiles in the liver and plasma of HFD‐fed mice

We used LC–MS/MS to elucidate the profiles of free fatty acids (FFAs) and metabolites of ω‐6 and ω‐3 PUFAs. Compared with the control diet, both HFD and ω3HFD increased the hepatic content of saturated fatty acid (Figure 2A). C18:2, C18:3 and C22:4 content was increased with the HFD and decreased with ω‐3 PUFA supplementation (Figure 2B). Hepatic ω‐3 PUFAs C20:5 (EPA) and C22:5 (DPA) content were significantly increased and that of ω‐6 PUFA C20:4 (ARA) was significantly decreased with ω3HFD compared to the HFD (Figure 2B). Therefore, ω‐6 content was lower and ω‐3 PUFA content was higher in mouse liver with ω3HFD than the HFD.

Figure 2.

Figure 2

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Alterations in hepatic FFA and PUFA metabolite (ω‐3 and ω‐6) profile determined by LC–MS/MS. Mice were fed a control diet (Ctrl), HFD or ω3HFD for 4 days. Hepatic (A) saturated fatty acid (FA) level, (B) unsaturated FA level and (C) eicosanoid profile of ω‐6 and ω‐3 PUFA metabolites shown by a heat map. (A–C) n = 6 animals per group. (D) Heat map showing eicosanoid profile of ω‐6 and ω‐3 PUFA metabolites in plasma samples. (E) Correlation network constructed with Pearson correlation coefficients (PCC >0.5 or <0.5). Positive and negative correlations are represented by violet and blue edges, respectively, and the darkness of the colour represents the degree of correlation. Green, blue, red and yellow dots represent EPA‐, DHA‐, ARA‐ and linoleic acid (LA)‐derived metabolites respectively. The dot size represents the connectiveness. (F) The plasma levels of ARA, EPA and DHA metabolites that were significantly changed by the HFD and reversed by ω3HFD. (D–F) n = 7 animals in the control group and n = 8 animals in HFD and ω3HFD groups (one sample in the control group was missing because of insufficient volume of plasma). In (A, B, F), the mean values of the control group were set to 1. The values of other groups were normalized to the control group values, presented as fold of control values. Data are mean ± SEM. *P < 0.05 versus Ctrl, #P < 0.05 versus HFD.

We explored targeted profiling of ARA and ω‐3 PUFAs (EPA and DHA) in liver (Figure 2C) and plasma (Figure 2D). In liver, ω‐3 PUFA supplementation significantly increased the contents of metabolites derived from EPA and DHA as compared with the HFD alone. Mouse liver with the control diet and ω‐3 PUFA supplementation showed similar increases in metabolite content (Figure S2F). Because eicosanoids in plasma have local and systemic biological actions, we measured the concentrations of ARA, EPA, DHA, docosapentaenoic acid (DPA) and their metabolites in plasma (Figure 2D and Table S2). Positive correlations for each eicosanoid pair (Figure 2E) are represented by violet edges and negative correlations by blue edges in Figure 2E. The darkness of the colour represents the degree of correlation. In general, the contents of ω‐3 PUFA‐derived metabolites were correlated with each other and those of EPA‐ and ARA‐derived metabolites were negatively correlated with each other directly. The dot size in the figure represents the extent of the correlation: the more connections for a node, the larger its size. No metabolites of ω‐3 PUFA showed a significantly higher correlation than others (Figure 2E). Additionally, in‐line with the degree of hepatic steatosis, the concentrations of the metabolites derived from EPA by lipoxygenase (15‐HEPE, 5‐HEPE, 9‐HEPE, 11‐HEPE and 12‐HEPE) and cytochrome P450 (18‐HEPE, 11,12‐EEQ, 8,9‐EEQ and 17,18‐EEQ) showed greater decrease with the HFD and increase with ω‐3 PUFA supplementation than those of other metabolites (Figure 2F). Thus, HEPEs and EEQs may play a critical role in the protective effects of ω‐3 PUFA on hepatic steatosis.

To investigate the direct effect of ω‐3 PUFAs and LC–MS/MS‐screened metabolites on the liver, primary hepatocytes were isolated and treated with palmitate with or without a mixture of EPA and DHA. However, ω‐3 PUFAs had no protective effect on palmitate‐induced lipid accumulation in primary hepatocytes (Figure S3A). Furthermore, the mixture of either their metabolite HEPEs (15‐HEPE, 5‐HEPE, 9‐HEPE, 11‐HEPE, 12‐HEPE and 18‐HEPE) or EEQs (11,12‐EEQ, 8,9‐EEQ, 14,15‐EEQ and 17,18‐EEQ) did not affect the steatosis in palmitate‐treated hepatocytes (Figure S3B). Thus, these data indicate the existence of an indirect protective effect of ω‐3 PUFAs on hepatic steatosis.

ω‐3 PUFA attenuated short‐term HFD‐induced adipose tissue inflammation

Inflammation reflected by macrophage infiltration in adipose tissue but not liver is a major phenotype after short‐term HFD treatment and is associated with hepatic lipid accumulation (Jung and Choi, 2014; Senthil Kumar et al., 2014). Indeed, the macrophage content in livers in our study was not induced by a short‐term HFD, as indicated by similar number of MAC‐3‐positive cells and the mRNA levels of CD68 and F4/80 in liver (Figure 3A, B). In contrast, the short‐term HFD significantly enhanced macrophage infiltration in adipose tissue, which was reversed by ω‐3 PUFA supplementation (Figure 3C, D). However, ω‐3 PUFA supplementation did not change the number of MAC‐3‐positive cells or mRNA levels of CD68 and F4/80 in adipose tissue with the control diet (Figure S2G, H). In parallel with the infiltration of adipose tissue macrophages (ATMs), the plasma content of C‐C motif chemokine ligand 2 (CCL2/monocyte chemoattractant protein 1), IL‐6 and TNFα was increased with the HFD and reduced by ω‐3 PUFA supplementation (Figure 3E). In addition, adipose tissue from ω3HFD‐fed mice attracted fewer macrophages than did tissue from HFD‐fed mice (Figure 3F). Therefore, ω‐3 PUFA can alleviate short‐term HFD‐induced inflammation in adipose tissue, which may improve hepatic steatosis by a crosstalk between adipose tissue and liver.

Figure 3.

Figure 3

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ω‐3 PUFA supplementation ameliorated short‐term HFD‐induced adipose inflammation in mice. Mice were fed a control diet (Ctrl), HFD or ω3HFD for 4 days. (A) Immunohistochemical analysis of MAC‐3 (for macrophages) in liver. (B) qPCR analysis of mRNA levels of markers of macrophages in liver. (C) Immunohistochemical analysis of MAC‐3 in adipose tissue. (D) qPCR analysis of mRNA levels of markers of macrophages in adipose tissue. (E) Levels of pro‐inflammatory cytokines in plasma (A–E, n = 8 animals per group). (F) Macrophage migration induced by adipose tissue (left panel) and quantification of macrophage migration by counting number of macrophages from DAPI staining (right panel), n = 5 animals per group. In (B) and (D), the mean values of the control group were set to 1. The values of other groups were normalized to control group values, represented as fold of control values. Data are mean ± SEM. *P < 0.05 versus Ctrl, #P < 0.05 versus HFD.

HEPEs and EEQs attenuated the inflammatory response in macrophages induced by palmitate

Macrophage infiltration and pro‐inflammatory cytokine secretion are considered new factors that regulate local adipose tissue inflammation and also contribute to liver diseases (Qureshi and Abrams, 2007; Jung and Choi, 2014). To investigate the effect of EPA and metabolites on macrophage inflammation, we treated peritoneal macrophages with palmitate to induce the inflammatory response. Consistent with previous studies (Mullen et al., 2010; De Boer et al., 2014), EPA at 50 μM had a significant protective effect on the expression of palmitate‐induced pro‐inflammatory cytokines in macrophages, including inducible NOS (iNOS), TNFα, IL‐1β and IL‐6 (Figure 4A). Next, we studied whether the anti‐inflammatory effect of EPA on macrophages was mediated by its metabolites based on the metabolomic results. HEPEs (a mixture of 5‐HEPE, 9‐HEPE, 11‐HEPE, 12‐HEPE, 15‐HEPE and 18‐HEPE with equal amounts of each) suppressed the mRNA levels of iNOS, TNFα, IL‐1β and IL‐6 induced by palmitate at 1 μM in macrophages (Figure 4B–E), whereas EEQs (a mixture of 8,9‐EEQ, 11,12‐EEQ, 14,15‐EEQ and 17,18‐EEQ with equal amounts of each) reduced the expression of iNOS at the same dose (Figure 4B). Furthermore, the anti‐inflammatory effect of HEPEs and EEQs was more pronounced than the same dose of EPA, except for suppression of iNOS expression (Figure 4B–E), so the metabolites may be more effective than EPA. The activation of MAPKs, including ERK, JNK and p38, mediates macrophage activation and pro‐inflammatory cytokine production (Rao, 2001). Consistent with a previous study, palmitate activated JNK and p38 rather than ERK in macrophages (Snodgrass et al., 2016). Only JNK activation induced by palmitate was reversed by both HEPEs and EEQs in macrophages (Figure 4F, G). Hence, HEPEs and EEQs, as metabolites of EPA, may mediate the anti‐inflammatory effect of EPA in macrophages to ameliorate adipose tissue inflammation.

Figure 4.

Figure 4

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HEPEs and EEQs derived from EPA attenuated macrophage inflammation. (A) Macrophages were treated with palmitate (PA) (200 μM) for 24 h to induce inflammation with or without 50 μM EPA. qPCR analysis of mRNA levels of iNOS, TNFα, IL‐1β and IL‐6. (B–E) Macrophages were treated with PA (200 μM) for 24 h; 1 μM EEQs (a mixture of 8,9‐EEQ, 11,12‐EEQ, 14,15‐EEQ and 17,18‐EEQ), HEPEs (a mixture of 5‐HEPE, 9‐HEPE, 11‐HEPE, 12‐HEPE, 15‐HEPE and 18‐HEPE) or EPA was added at the same time as PA. qPCR analysis of mRNA levels of iNOS (B), TNFα (C), IL‐1β (D) and IL‐6 (E). (F, G) Macrophages were treated with PA (200 μM) for 8 h and 1 μM EEQs, HEPEs or EPA was added at the same time as PA. Western blot analysis of protein levels of p‐JNK, JNK, p‐p38, p38, p‐ERK and ERK. n = 5 independent experiments. The mean values of the control diet (Ctrl) group were set to 1. The values of other groups were normalized to control group values, represented as fold of control values. Data are mean ± SEM. *P < 0.05 versus Ctrl, #P < 0.05 versus PA.

17,18‐EEQ, 5‐HEPE and 9‐HEPE attenuated the inflammatory response in macrophages treated with palmitate

To identify the effective components of HEPEs and EEQs, we further studied the function of each EEQ (8,9‐EEQ, 11,12‐EEQ, 14,15‐EEQ or 17,18‐EEQ) and HEPE (5‐HEPE, 9‐HEPE, 11‐HEPE, 12‐HEPE, 15‐HEPE or 18‐HEPE) in macrophages. Among four EEQs and six HEPEs, 17,18‐EEQ greatly decreased TNFα and IL‐6 expression in palmitate‐treated macrophages; 9‐HEPE suppressed iNOS, TNFα, IL‐1β and IL‐6 expression, and 5‐HEPE suppressed only iNOS expression (Figure 5A–D). In addition, 17,18‐EEQ significantly decreased CCL3 and CCL4 expression; 5‐HEPE decreased CCL2 and CCL4 expression; and 9‐HEPE significantly decreased CCL2, CCL3, CCL4 and CCL7 expression (Figure 5E). Correspondingly, macrophage migration was increased by pretreatment with palmitate, which was significantly reduced by the treatment with 17,18‐EEQ, 5‐HEPE and 9‐HEPE (Figure 5F). In addition, consistent with the effect of EEQ and HEPE mixtures on MAPK signalling, 17,18‐EEQ, 5‐HEPE and 9‐HEPE significantly suppressed palmitate‐induced JNK but not p38 activation (Figure 5G, H). In contrast, other EEQs and HEPEs had little effect on the expression of palmitate‐induced pro‐inflammatory cytokines (Figure S4). Thus, 17,18‐EEQ, 5‐HEPE and 9‐HEPE were more effective than other HEPEs and EEQs on suppressing the palmitate‐induced inflammatory response in macrophages.

Figure 5.

Figure 5

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17,18‐EEQ (17,18E), 9‐HEPE (9H) and 5‐HEPE (5H) derived from EPA attenuated macrophage inflammation induced by palmitate (PA). (A–F) Macrophages were treated with PA (200 μM) for 24 h to induce inflammation. 17,18E (1 μM), 5H (1 μM) or 9H (1 μM) was added at the same time as PA treatment. qPCR analysis of mRNA levels of iNOS (A), TNFα (B), IL‐1β (C), IL‐6 (D) and chemokines including CCL2, CCL3, CCL4, CCL5 and CCL7 (E). (F) Macrophage migration induced by PA treatment with or without 17,18E, 5H or 9H (left panel) and quantification of macrophage migration by counting number of macrophages from DAPI staining (right panel). (G, H) Macrophages were treated with PA (200 μM) for 8 h, and 17,18E (1 μM), 5H (1 μM) or 9H (1 μM) was added at the same time as PA treatment. Western blot analysis of protein levels of p‐JNK, JNK, p‐p38, p38, p‐ERK and ERK. In (A–E, H), the mean values of the control (Ctrl) group were set to 1. The values of other groups were normalized to the control group values, presented as fold of control values. n = 5 independent experiments. Data are mean ± SEM. *P < 0.05 versus Ctrl, #P < 0.05 versus PA.

A mixture of 17,18‐EEQ, 5‐HEPE and 9‐HEPE ameliorated short‐term HFD‐induced hepatic steatosis and adipose tissue inflammation

To elucidate the effects of 17,18‐EEQ, 5‐HEPE and 9‐HEPE on short‐term HFD‐induced hepatic steatosis in vivo, we injected, i.p., a mixture of equal amounts of 17,18‐EEQ, 5‐HEPE and 9‐HEPE into mice fed the control diet and HFD. Daily food intake was not changed by the mixture added to the control diet or HFD (Figure S5), nor did it change hepatic lipid content, plasma total cholesterol or LDL‐cholesterol level or the number of macrophages in adipose tissue of mice fed the control diet (Figure 6A–G). In contrast, the mixture greatly ameliorated short‐term HFD‐induced hepatic steatosis, with intact morphology (Figure 6A). Hepatic TG rather than total cholesterol level was significantly decreased (Figure 6B, C), and plasma total cholesterol and LDL‐cholesterol levels were attenuated by the mixture in HFD‐fed mice (Figure 6D, E). The mixture also suppressed short‐term HFD‐induced adipose tissue inflammation, as evidenced by reduced number of MAC‐3‐positive cells and mRNA levels of macrophage markers CD68 and F4/80 (Figure 6F, G). Therefore, the protective effect of ω‐3 PUFA on short‐term HFD‐induced hepatic steatosis was mediated by 17,18‐EEQ, 5‐HEPE and 9‐HEPE in macrophages, which attenuated the inflammation in adipose tissue (Figure 7).

Figure 6.

Figure 6

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A mixture of 17,18‐EEQ, 9‐HEPE and 5‐HEPE inhibited short‐term HFD‐induced hepatic steatosis and adipose inflammation. Mice were fed the control diet (Ctrl) or HFD diet with or without an i.p. injection of a mixture of 17,18‐EEQ, 9‐HEPE and 5‐HEPE. (A) Oil red O staining (ORO) (upper panel) and haematoxylin and eosin (H&E) staining (lower panel). (B, C) Quantitative assay of TG (B) and cholesterol (CHO) (C) content in liver. (D, E) Plasma CHO (D) and LDL‐CHO (LDL‐C) (E) level. (F) Immunohistochemical analysis of MAC‐3 level in adipose tissue. (G) qPCR analysis of levels of macrophage markers in adipose tissue and the mean values for the control group were set to 1. The values of other groups were normalized to the control group values, presented as fold of control values. HFD + Mix, HFD plus the mixture of 17,18‐EEQ, 9‐HEPE and 5‐HEPE; n = 8 animals per group. *P < 0.05 versus Ctrl, #P < 0.05 versus HFD.

Figure 7.

Figure 7

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Proposed mechanism of ω‐3 PUFA mediated amelioration of short‐term HFD‐induced adipose tissue inflammation and fatty liver.

Discussion

ω‐3 PUFAs (EPA and DHA) are considered to have a protective role in metabolic disorders. In our study, ω‐3 PUFA prevented the onset of NAFLD induced by acute HFD in mice, which was mediated by 17,18‐EEQ, 5‐HEPE and 9‐HEPE by inhibiting the inflammatory response in adipose tissue macrophages.

Our and other studies showed that short‐term HFD exposure increased insulin resistance, plasma LDL‐cholesterol levels, hepatic TG levels and macrophage inflammatory response in adipose tissue rather than liver (Lee et al., 2011; Senthil Kumar et al., 2014). In‐line with other findings (Senthil Kumar et al., 2014), we found decreased ACC expression in the liver of short‐term HFD‐fed mice, which may be caused by a negative feedback effect of lipid overload. Additionally, ω‐3 PUFA supplementation further reduced the expression of FAS and ACC and increased that of genes involved in β‐oxidation. Intriguingly, the content of C16:0 was increased with both the HFD and ω3HFD compared with the control diet, despite their lower expression of ACC than controls. This finding may be due to the abundant palmitate in the food, which is absorbed by the small intestine and stored in adipose tissue as TG. Adipose tissue then releases FFAs in the circulation by lipolysis. The lipolysis of stored TG in adipocytes and dietary fat are the main sources of circulating FFAs, which can be taken up by the liver and are another important source of FFAs in liver besides de novo lipogenesis (Bradbury, 2006). In addition, although ω3HFD supplementation decreased ACC expression to a greater degree than did the HFD, the expression of CD36 was increased; CD36 mediates lipid uptake and might contribute to the comparable contents of C16:0 between the HFD and ω3HFD groups. The improvement in hepatic lipid accumulation by ω‐3 PUFA may result from reduced de novo lipogenesis and increased lipid β‐oxidation.

ω‐3 PUFA supplementation alters the metabolic profile of ω‐3 and ω‐6 types, and the change is extensive, multiple and complicated (Zhang et al., 2015). LC–MS/MS–based targeted metabolomics is the only effective strategy to study the levels of metabolites in NAFLD. In our previous study, we quantified eicosanoids in mouse and human plasma and mouse aorta samples, after ω‐3 PUFA supplementation, by LC–MS/MS (Zhang et al., 2015). Here, we found that the levels of several HEPEs and EEQs derived from EPA were decreased in HFD mouse plasma and markedly increased by ω‐3 PUFA supplementation. Thus, HEPEs and EEQs may play an important role in short‐term HFD‐induced NAFLD.

Several studies have investigated the function of HEPEs in various diseases. For instance, 5‐HEPE enhanced glucose‐dependent insulin secretion in pancreatic beta cells by activating the GPR119/cAMP pathway (Kogure et al., 2011). 18‐HEPE injection prevented pressure overload‐induced adaptive cardiac remodelling via its anti‐inflammatory effects (Endo et al., 2014). Also, a recent study revealed the protective effects of EEQs on NAFLD. 17,18‐EEQ and 19,20‐epoxydocosapentaenoic acid were shown to restore autophagy and reduce ER stress in palmitate‐treated hepatocytes and adipocytes (Lopez‐Vicario et al., 2015). As a result, a soluble epoxide hydrolase inhibitor further reduced HFD‐induced hepatic steatosis in fat‐1 mice because of the increased hepatic 17,18‐EEQ and 19,20‐EDP levels (Lopez‐Vicario et al., 2015). GPR119 activated by oleoylethanolamide can suppress food intake and reduce body weight gain and white adipose tissue accumulation in high‐fat‐fed rats (Overton et al., 2006). However, the food intake was not affected by ω‐3 PUFA supplementation in our study. Therefore, we studied the direct effect of HEPEs and EEQs on hepatocytes. Unexpectedly, the mixture of EPA/DHA, HEPEs or EEQs did not attenuate palmitate‐induced lipid accumulation in primary hepatocytes, which implied an indirect effect of ω‐3 PUFA and metabolites on liver.

Adipose tissue inflammation can be induced by HFD before hepatic inflammation occurs (Senthil Kumar et al., 2014). It is demonstrated as increased macrophage infiltration, altered macrophage polarization and increased pro‐inflammatory cytokines in plasma (Lee et al., 2011). Inflammatory factors produced by ATMs are considered to contribute to the crosstalk between local adipose tissue inflammation and hepatic steatosis (Tilg, 2010; Tilg and Moschen, 2010). For example, TNFα can induce hepatic steatosis in mice (Endo et al., 2007). IL‐1β secreted from adipose tissue can regulate fat–liver crosstalk in obesity (Nov et al., 2013). IL‐6 administration can elevate fat liver content and aggravated steatosis in HFD‐induced obese mice despite the expression of lipogenic genes being suppressed (Gavito et al., 2016). A clinical trial found that ω‐3 PUFA supplementation reduced the expression of IL‐6, IL‐8, TNFα and toll‐like receptor 4 in adipose tissue of obese pregnant women (Haghiac et al., 2015). As one of the most important components of ω‐3 PUFA, EPA was demonstrated to reduce adipocyte hypertrophy and inflammation in long‐term HFD‐induced obese mice (Kalupahana et al., 2010; LeMieux et al., 2015). In our study, ω‐3 PUFA significantly ameliorated adipose tissue inflammation induced by short‐term HFD and decreased pro‐inflammatory cytokine production. Pro‐inflammatory cytokines released from ATMs are believed to be the main source of adipose tissue inflammation (Qureshi and Abrams, 2007; Jung and Choi, 2014). EPA and DHA have anti‐inflammatory effects by targeting macrophages, which is helpful to attenuate several diseases. EPA and DHA can repress LPS‐induced inflammatory cytokine secretion and activation of the NF‐κB pathway in THP‐1‐derived macrophages (Mullen et al., 2010; Snodgrass et al., 2016). DHA can ameliorate the inflammatory response in macrophages and improve insulin sensitivity in adipocytes (Rao, 2001). Furthermore, EPA was more effective than DHA at inhibiting LPS‐induced inflammatory effects in asthmatic alveolar macrophages (Mickleborough et al., 2009). However, the effective components of the metabolites that mediate the anti‐inflammatory effect of EPA/DHA are still unknown.

To answer this question, we next investigated the function of HEPEs and EEQs in macrophages. We used palmitate, whose level is systemically elevated in diet‐induced obesity, to induce an inflammatory response in macrophages. HEPEs and EEQs suppressed the expression of pro‐inflammatory cytokines and JNK activation induced by palmitate in macrophages. Furthermore, 17,18‐EEQ, 5‐HEPE and 9‐HEPE were more efficient than other EEQs and HEPEs in inhibiting the inflammatory response in macrophages. In‐line with these findings, the mixture of 17,18‐EEQ, 5‐HEPE and 9‐HEPE significantly ameliorated short‐term HFD‐induced lipid accumulation in the liver and adipose tissue inflammation. These results revealed the anti‐steatotic effect of 17,18‐EEQ, 5‐HEPE and 9‐HEPE, which contributed to the suppression of the inflammatory response in ATMs.

In summary, ω‐3 PUFA ameliorates the onset of NAFLD induced by short‐term HFD. From the metabolomics, 17,18‐EEQ, 5‐HEPE and 9‐HEPE were identified to attenuate the inflammatory response of macrophages via the JNK pathway, which ameliorated adipose tissue inflammation and protected against hepatic steatosis induced by a short‐term HFD. 17,18‐EEQ, 5‐HEPE and 9‐HEPE may be more effective than ω‐3 PUFA at preventing NAFLD, at the early stage, in people with an excessive intake of fat.

Author contributions

C.W. and W.L. contributed to the concept and design, data acquisition, analysis and interpretation and drafting of the article. L.Y. contributed to data acquisition, analysis and interpretation and drafting of the article. X.Z., X.Z., C.Y. and J.H. contributed to data acquisition of the article. H.J. and Y.Z. contributed to the concept and design of the article. D.A. contributed to the concept and design, data acquisition, analysis and interpretation of data of the article. D.A. is the guarantor of the work.

Conflict of interest

The authors declare no conflicts of interest.

Declaration of transparency and scientific rigour

This Declaration acknowledges that this paper adheres to the principles for transparent reporting and scientific rigour of preclinical research recommended by funding agencies, publishers and other organisations engaged with supporting research.

Supporting information

 

Click here for additional data file. (1.2MB, pdf)

Table S1 List of oligonucleotide primer pairs used in qPCR.

Table S2 The plasma level of ARA, EPA, DHA, DPA and metabolites.

Click here for additional data file. (190KB, pdf)

Figure S1 Short‐term HFD‐induced glucose intolerance and tissue/body wt not changed by ω‐3 PUFA supplementation. Mice were fed control, HFD or ω‐3 PUFA‐enriched HFD (ω3HFD) for 4 days. (A) Daily food intake of each mouse. (B) Intraperitoneal glucose tolerance testing (IPGTT) was performed, and glucose levels were measured. (C) The body wt and (D) liver, adipose tissue and heart wt. (E) Ratios of liver to body wt, adipose tissue to body wt and heart to body wt. n = 8 animals per group, data are mean±SEM. *P < 0.05 versus Ctrl.

Figure S2 The increased ω‐3 PUFA derived metabolites by ω‐3 PUFA supplementation in control diet had no effect on metabolic conditions in liver and macrophage amounts in adipose tissue. Mice were treated with control diet with or without ω‐3 PUFA supplementation (3%, wt/wt). (A) Oil‐red O staining (ORO) (upper panel) and H&E staining (lower panel) of liver. (B‐C) Quantitative assay of triglycerides (TG) content (B) and cholestrol (CHO) content (C) in liver. (D‐E) Plasma cholesterol (D) and LDL‐cholesterol (LDL‐C) (E) levels. (F) Eicosanoid profile with of ω‐6 and ω‐3 PUFA metabolitsmetabolites in liver (F). (G) Immunohistochemical analysis of MAC‐3 in adipose tissue. (H) qPCR analysis of mRNA levels of markers of macrophages in adipose tissue. The mean values of the control group were set to 1. The values of other groups were normalized to control group values, represented as fold of control values. n = 7 animals per group, data are mean±SEM.

Figure S3 ω‐3 PUFAs had no direct protective effect on palmitate‐treated primary hepatocyte. (A‐B) Primary hepatocytes were treated with palmitate (100 μM) for 24 hr to induce lipid accumulation. The mixture of EPA and DHA (1:1, total amount 50 μM) (A), EEQs (1 μM) or HEPEs (1 μM) (B) was added at the same time as palmitate. ORO staining was performed in hepatocytes (left panel) and the dye in hepatocyte was dissolved and analysed at emission 570 nm (right panel). Ctrl, control; PA, palmitate; PA+(E/D), palmitate with the mixture of EPA and DHA; PA + EEQs, palmitate with the mixture of EEQs; PA + HEPEs, palmitate with the mixture of HEPEs. n = 5 independent experiments. The mean values of the control group were set to 1. The values of other groups were normalized to control group values, represented as fold of control values. Data are mean±SEM. *P < 0.05 vs Ctrl.

Figure S4 8,9‐EEQ, 11,12‐EEQ, 14,15‐EEQ, 11‐HEPE, 12‐HPEP, 15‐HEPE or 18‐HEPE had little effect on macrophage inflammation induced by palmitate. Macrophages were treated with palmitate (200 μM) for 24 hr to induce inflammation. 8,9‐EEQ, 11,12‐EEQ, 14,15‐EEQ, 11‐HEPE, 12‐HPEP, 15‐HEPE or 18‐HEPE (all 1 μM) was added at the same time as palmitate treatment. (A‐D) qPCR analysis of mRNA levels of iNOS (A), TNFα (B), IL‐1β (C) and IL‐6 (D). Ctrl, control; PA, palmitate; 8,9E, 8,9‐EEQ; 11,12E, 11,12‐EEQ; 14,15E, 14,15‐EEQ; 11H, 11‐HEPE; 12H, 12‐HPEP; 15H, 15‐HEPE; 18H, 18‐HEPE. n = 5 independent experiments. The mean values of the control group were set to 1. The values of other groups were normalized to control group values, represented as fold of control values. Data are mean±SEM. *P < 0.05 Ctrl, # P < 0.05 vs PA.

Figure S5 Mixture of 17,18‐EEQ, 9‐HEPE and 5‐HEPE did not change the food intake compared with HFD group. Mice were fed the control or HFD diet with or without intraperitoneal injection of a mixture of 17,18‐EEQ, 9‐HEPE and 5‐HEPE. (A) Daily food intake of each mouse. HFD + Mix, high fat diet plus the mixture of 17,18‐EEQ, 9‐HEPE and 5‐HEPE; n = 8 animals per group.

Click here for additional data file. (252.9KB, pdf)

Acknowledgements

This work was supported by the National Natural Science Foundation of China (81500445, 81420108003 and 91539108) and Tianjin Educational Committee Foundation (no. 20140102).

Wang, C. , Liu, W. , Yao, L. , Zhang, X. , Zhang, X. , Ye, C. , Jiang, H. , He, J. , Zhu, Y. , and Ai, D. (2017) Hydroxyeicosapentaenoic acids and epoxyeicosatetraenoic acids attenuate early occurrence of nonalcoholic fatty liver disease. British Journal of Pharmacology, 174: 2358–2372. doi: 10.1111/bph.13844.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

 

Click here for additional data file. (1.2MB, pdf)

Table S1 List of oligonucleotide primer pairs used in qPCR.

Table S2 The plasma level of ARA, EPA, DHA, DPA and metabolites.

Click here for additional data file. (190KB, pdf)

Figure S1 Short‐term HFD‐induced glucose intolerance and tissue/body wt not changed by ω‐3 PUFA supplementation. Mice were fed control, HFD or ω‐3 PUFA‐enriched HFD (ω3HFD) for 4 days. (A) Daily food intake of each mouse. (B) Intraperitoneal glucose tolerance testing (IPGTT) was performed, and glucose levels were measured. (C) The body wt and (D) liver, adipose tissue and heart wt. (E) Ratios of liver to body wt, adipose tissue to body wt and heart to body wt. n = 8 animals per group, data are mean±SEM. *P < 0.05 versus Ctrl.

Figure S2 The increased ω‐3 PUFA derived metabolites by ω‐3 PUFA supplementation in control diet had no effect on metabolic conditions in liver and macrophage amounts in adipose tissue. Mice were treated with control diet with or without ω‐3 PUFA supplementation (3%, wt/wt). (A) Oil‐red O staining (ORO) (upper panel) and H&E staining (lower panel) of liver. (B‐C) Quantitative assay of triglycerides (TG) content (B) and cholestrol (CHO) content (C) in liver. (D‐E) Plasma cholesterol (D) and LDL‐cholesterol (LDL‐C) (E) levels. (F) Eicosanoid profile with of ω‐6 and ω‐3 PUFA metabolitsmetabolites in liver (F). (G) Immunohistochemical analysis of MAC‐3 in adipose tissue. (H) qPCR analysis of mRNA levels of markers of macrophages in adipose tissue. The mean values of the control group were set to 1. The values of other groups were normalized to control group values, represented as fold of control values. n = 7 animals per group, data are mean±SEM.

Figure S3 ω‐3 PUFAs had no direct protective effect on palmitate‐treated primary hepatocyte. (A‐B) Primary hepatocytes were treated with palmitate (100 μM) for 24 hr to induce lipid accumulation. The mixture of EPA and DHA (1:1, total amount 50 μM) (A), EEQs (1 μM) or HEPEs (1 μM) (B) was added at the same time as palmitate. ORO staining was performed in hepatocytes (left panel) and the dye in hepatocyte was dissolved and analysed at emission 570 nm (right panel). Ctrl, control; PA, palmitate; PA+(E/D), palmitate with the mixture of EPA and DHA; PA + EEQs, palmitate with the mixture of EEQs; PA + HEPEs, palmitate with the mixture of HEPEs. n = 5 independent experiments. The mean values of the control group were set to 1. The values of other groups were normalized to control group values, represented as fold of control values. Data are mean±SEM. *P < 0.05 vs Ctrl.

Figure S4 8,9‐EEQ, 11,12‐EEQ, 14,15‐EEQ, 11‐HEPE, 12‐HPEP, 15‐HEPE or 18‐HEPE had little effect on macrophage inflammation induced by palmitate. Macrophages were treated with palmitate (200 μM) for 24 hr to induce inflammation. 8,9‐EEQ, 11,12‐EEQ, 14,15‐EEQ, 11‐HEPE, 12‐HPEP, 15‐HEPE or 18‐HEPE (all 1 μM) was added at the same time as palmitate treatment. (A‐D) qPCR analysis of mRNA levels of iNOS (A), TNFα (B), IL‐1β (C) and IL‐6 (D). Ctrl, control; PA, palmitate; 8,9E, 8,9‐EEQ; 11,12E, 11,12‐EEQ; 14,15E, 14,15‐EEQ; 11H, 11‐HEPE; 12H, 12‐HPEP; 15H, 15‐HEPE; 18H, 18‐HEPE. n = 5 independent experiments. The mean values of the control group were set to 1. The values of other groups were normalized to control group values, represented as fold of control values. Data are mean±SEM. *P < 0.05 Ctrl, # P < 0.05 vs PA.

Figure S5 Mixture of 17,18‐EEQ, 9‐HEPE and 5‐HEPE did not change the food intake compared with HFD group. Mice were fed the control or HFD diet with or without intraperitoneal injection of a mixture of 17,18‐EEQ, 9‐HEPE and 5‐HEPE. (A) Daily food intake of each mouse. HFD + Mix, high fat diet plus the mixture of 17,18‐EEQ, 9‐HEPE and 5‐HEPE; n = 8 animals per group.

Click here for additional data file. (252.9KB, pdf)

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