Dual action of eicosapentaenoic acid in hepatoma cells: up-regulation of metabolic action of insulin and inhibition of cell proliferation.

Exogenous administration of eicosapentaenoic acid (EPA) improves insulin sensitivity, but its precise mechanism remains unknown. Here we show that EPA stimulates the intracellular insulin signaling pathway in hepatoma cells. Exposure of these cells to EPA caused up-regulation of several insulin-induced activities including tyrosine phosphorylation of insulin receptor substrate-1, insulin receptor substrate-1-associated phosphatidylinositol 3-kinase, and its downstream target Akt kinase activity as well as down-regulation of gluconeogenesis. In contrast, EPA decreased mitogen-activated protein kinase activity and inhibited cell proliferation. These findings raise the possibility that EPA up-regulates metabolic action of insulin and inhibits cell growth in humans.

anti-IRS-1 for 1 h and then horseradish peroxidase-conjugated antirabbit IgG for 1 h to detect association of GRB2 or PI3K with IRS-1. A chemiluminescent peroxidase substrate (ECL; Amersham Pharmacia Biotech) was applied according to the manufacturer's instructions, and the membranes were exposed briefly to x-ray film.
MAPK in Vitro Kinase Assays-MAPK activity of HepG2 cells was measured by using MAPK assay kit according to the manufacturer's instructions. In short, the cells were washed with PBS, lysed in lysis buffer (20 mM Tris (pH 7.5), 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1% Triton X-100, 2.5 mM sodium pyrophosphate, 1 mM ␤-glycerophosphate, 1 mM Na 3 VO 4 , 1 l/ml leupeptin, 1 mM phenylmethylsulfonyl fluoride), scraped off the dishes, transferred to microcentrifuge tubes, sonicated, and microcentrifuged for 10 min at 4°C. The supernatant was used for immunoprecipitation of activated MAPKs. The supernatant was incubated with immobilized phospho-p44/42 MAPK (Thr 202 /Tyr 204 ) monoclonal antibody for 4 h and microcentrifuged for 30 s at 4°C. The pellets were washed twice with ice-cold lysis buffer and twice with kinase buffer (25 mM Tris (pH 7.5), 5 mM ␤-glycerophosphate, 2 mM dithiothreitol, 0.1 mM Na 3 VO 4 , 10 mM MgCl 2 ). The pellets were incubated with 200 M ATP and 2 g of Elk-1 fusion protein for 30 min at 30°C. The reaction was terminated by 25 l of 3ϫ SDS sample buffer (187.5 mM Tris-HCl (pH 6.8), 6% (w/v) SDS, 30% glycerol, 150 mM dithiothreitol, 0.3% (w/v) bromphenol blue). Samples were boiled, separated by electrophoresis through a 10% SDS-polyacrylamide gel, and transferred to polyvinylidene difluoride membrane for 30 min. Immunoblot was blocked by incubation in TBST (10 mM Tris-HCl, pH 7.6, 150 mM NaCl, 0.1% Tween 20) containing 5% skim milk at room temperature for 1 h and then probed with a 1:1000 dilution of anti-phospho-Elk-1 antibody. The membranes were washed three times in TBST and incubated with a 1:1000 dilution of horseradish peroxidase-linked anti-rabbit IgG antibody and then reacted with LumiGLO reagent according to the manufacturer's instructions. The membranes were exposed briefly to x-ray film.
Akt in Vitro Kinase Assays-Akt kinase activity of HepG2 cells was measured by using Akt kinase assay kit according to the manufacturer's instructions. In short, the cells were washed with PBS, lysed in lysis buffer (20 mM Tris (pH 7.5), 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1% Triton X-100, 2.5 mM sodium pyrophosphate, 1 mM ␤-glycerophosphate, 1 mM Na 3 VO 4 , 1 l/ml leupeptin, 1 mM phenylmethylsulfonyl fluoride), scraped off the dishes, transferred to microcentrifuge tubes, sonicated, and microcentrifuged for 10 min at 4°C. The supernatant was used for immunoprecipitation of activated Akt kinase. The supernatant was incubated with immobilized Akt antibody for 2-3 h and microcentrifuged for 30 s at 4°C. The pellets were washed twice with ice-cold lysis buffer and twice with kinase buffer (25 mM Tris (pH 7.5), 5 mM ␤-glycerophosphate, 2 mM dithiothreitol, 0.1 mM Na 3 VO 4 , 10 mM MgCl 2 ). The pellets were incubated with 200 M ATP and 1 g of GSK-3 fusion protein for 30 min at 30°C. The reaction was terminated by 25 l of 3ϫ SDS sample buffer (187.5 mM Tris-HCl (pH 6.8), 6% (w/v) SDS, 30% glycerol, 150 mM dithiothreitol, 0.03% (w/v) bromphenol blue). Samples were boiled, separated by electrophoresis through a 12% SDS-polyacrylamide gel, and transferred to polyvinylidene difluoride membrane for 30 min. Immunoblot was blocked by incubation in TBST (10 mM Tris-HCl, pH 7.6, 150 mM NaCl, 0.1% Tween 20) containing 5% skim milk at room temperature for 1 h and then probed with a 1:1000 dilution of anti-phospho-GSK-3 antibody. The membranes were washed three times in TBST and incubated with a 1:1000 dilution of horseradish peroxidase-linked anti-rabbit IgG antibody and then reacted with LumiGLO reagent according to the manufacturer's instructions. The membranes were exposed briefly to x-ray film.
Reverse Transcriptase-Polymerase Chain Reaction-Total RNA was prepared from H4-II-E cells using Trizol according to the manufacturer's instructions (Life Technologies, Inc.). One g total RNA was reverse-transcribed for 1 h at 37°C in 20 l of reaction medium made up of 50 mM Tris/HCl, 75 mM KCl, 3 mM MgCl 2 , 10 mM dithiothreitol, pH 8.3, 1 mM dNTPs, 15 pmol of antisense primer, and 200 units Moloney murine leukemia virus reverse transcriptase (Promega). The PCR amplifications were performed in 50 l of medium containing 50 mM Tris-HCl, 50 mM KCl, 1 mM MgCl 2 , pH 9.0, 0.2 mM dNTPs, 15 pmol of sense and antisense primers, and 2.5 units of Taq DNA polymerase (Promega). The reaction mixtures were subjected to 40 cycles of PCR amplification of PEPCK cDNA consisting of denaturation for 60 s at 94°C, annealing for 60 s at 56°C, and elongation for 120 s at 72°C. The oligonucleotide primers (5Ј-AGCCTCGACAGCCTGCCCCAGG-3Ј sense and 5Ј-CCAGTTGTTGACCAAAGGCTTTT-3Ј antisense) allowed amplification of a 575-bp PEPCK cDNA product (17) and verified with DNA sequencing (model 310, PerkinElmer Life Sciences). The ␤-actin sequence of the upstream primer was 5Ј-TGACCCAGATCATGTTT-GAGAGACC-3Ј, and the sequence of the downstream primer was 5Ј-CCATACCCAAGAAGGAAGGC-3Ј. After an initial denaturation for 30 s at 94°C, PCR was performed for 40 cycles. The conditions for PCR were 94°C for 60 s, 60°C for 60 s, and 72°C for 60 s. The rat ␤-actincoding region was obtained by reverse transcriptase-PCR and verified with DNA sequencing.
Northern Blot Analysis-Total RNA was prepared from cultured H4-II-E cells using Trizol. Twenty g of total RNA was heat-denatured for 10 min at 70°C in buffer containing 50% formamide, 2.4 M formaldehide, 1 ϫ 0.02 M MOPS, 50 mM sodium acetate, 10 mM Na 2 EDTA, and then loaded onto a 1.2% agarose/formaldehide gel made up in 1ϫ MOPS, pH 7.4, at 50 V for 3-4 h. After electrophoresis, RNA was transferred onto a Hybond N membrane by capillary action and fixed onto the filters by ultraviolet light cross-linking. The blot was probed with either a rat PEPCK or rat ␤-actin probe labeled by the Gene Images random prime labeling module and detected by the Gene Images CDP-star detection module according to the manufacturer's instructions, and the membranes were exposed to x-ray film for 30 -60 min.
Cell Proliferation Assays-Approximately 48 -72 h before the assay, cells were seeded into 24-well plates at 1-3 ϫ 10 5 cells/well. When cells reached 60 -70% confluence, the medium was replaced with Dulbecco's modified Eagle's medium containing 0.1% BSA, and then EPA, docosahexaenoic acid, arachidonic acid, oleic acid, or palmitic acid was added at the indicated concentration. The cultures were incubated in a CO 2 incubator for 72 h at 37°C unless otherwise indicated. Cells were washed with phosphate-buffered saline and stripped with 0.05% trypsin, 0.53 mM EDTA. The cell number was then determined using a cell counter. All samples were assayed in duplicate, and each experiment was repeated at least three times. Representative data are presented.
Cell proliferation assays were also performed by CellTiter 96 AQueous One Solution Cell Proliferation Assay kit according to the supplier (Life Technologies). Briefly, cells were seeded into 96-well plates and treated as described under cell proliferation assay. The cultures were incubated in a CO 2 incubator for 72 h at 37°C and then added to 20 l/well of CellTiter 96 AQueous One Solution Reagent, which contains a tetrazolium compound, MTS was added to the culture medium. The MTS tetrazolium compound is bioreduced by cells into a colored formazan product that is soluble in tissue culture medium. This conversion is presumably accomplished by NADPH or NADH produced by dehydrogenase enzymes in metabolically active cells (18). After incubating the plate 2-4 h in a CO 2 incubator, the absorbance at 490 nm was recorded with a 96-well plate reader. All samples were assayed in duplicate, and each experiment was repeated at least three times.
Cell Death Assays-Cells were treated as described under "Cell Proliferation Assays." Live and dead cells from each subculture were counted by trypan blue at sequential time points during incubation, and the ratio of dead/(dead ϩ live) cells was calculated. The mean values of these ratios and the S.E. were calculated at each time point.
Statistical Analysis-The data were expressed as the mean Ϯ S.E. unless noted otherwise. Statistics were analyzed by one-way repeated measures analysis of variance with a significant level of 0.05.

RESULTS
The effect of EPA on the tyrosine-phosphorylated cellular proteins was studied in HepG2 cells by immunoblot analysis with antibodies to phosphotyrosine (anti-Tyr(P)). Treatment for 24 h with 100 M EPA caused a significant increase in tyrosine-phosphorylated IRS-1 levels in HepG2 cells compared with those with vehicle treatment (455 Ϯ 44 versus 105 Ϯ 6% basal level, p Ͻ 0.01, n ϭ 5) (Fig. 1A). When HepG2 cells were incubated with 1-1000 M EPA for 24 h, EPA caused a significant and dose-dependent increase in tyrosine-phosphorylated IRS-1 levels in the cells as shown in Fig. 1B. HepG2 cells were treated with 100 M EPA for 24 h, followed by stimulation with 100 nM insulin for 1 min. EPA and insulin significantly increased tyrosine-phosphorylated IRS-1 levels up to 362 Ϯ 34 and 616 Ϯ 35 densitometric units (DU), respectively, compared with vehicle alone (73 Ϯ 6 DU). Combined stimulation with EPA and insulin resulted in a further increase of tyrosinephosphorylated IRS-1 levels up to 714 Ϯ 34 DU (p Ͻ 0.01, n ϭ 5) (Fig. 1C). The effect of EPA on IRS-1 phosphorylation is not mediated via insulin receptor (IR), because tyrosine phosphorylation of the insulin receptor ␤ chain was not different be-tween the presence and absence of EPA (Fig. 1D). EPA also increased tyrosine phosphorylation of IRS-1 in H4-II-E cells ( Fig. 2A). Furthermore, we tested the effect of other n-3 PUFAs, docosahexaenoic acid, n-6 PUFAs, arachidonic acid, monounsaturated fatty acid, oleic acid, saturated fatty acid, and palmitic acid on the tyrosine phosphorylation of IRS-1. Other fatty acids, except EPA, did not increase the tyrosine phosphorylation of IRS-1 (Fig. 2B). In addition, acute incubation of HepG2 cells with 100 M EPA for 0 -30 min did not increase in tyrosine-phosphorylated IRS-1 levels (data not shown). We investigated the effect of EPA on tyrosine phosphorylation of IRS-1 in other cell lines, rat skeletal muscle cells, and L6 myocytes. EPA did not increase tyrosine phosphorylation of IRS-1 in L6 cells (data not shown).
To investigate whether the effect of EPA on tyrosine phosphorylation is specific for IRS-1, we tested the effect of EPA on tyrosine phosphorylation of IRS-2 and SHC. HepG2 cells were treated with 100 M EPA for 24 h, followed by stimulation with 100 nM insulin for 1 min. EPA did not increase tyrosine phosphorylation of IRS-2 (Fig. 2C) (Fig. 2D).
Downstream signaling of IRS-1 is mediated by several associated proteins, including GRB2 and PI3K (19). We therefore tested the effect of EPA on the interaction of GRB2 or PI3K with IRS-1. When HepG2 cells were pretreated with 100 M EPA for 24 h, followed by stimulation with 100 nM insulin for 3 min, the amount of GRB2 associated IRS-1 (basal, 118 Ϯ 16 DU) was increased either by EPA (214 Ϯ 26 DU, p Ͻ 0.05) or by insulin (298 Ϯ 37 DU, p Ͻ 0.01, n ϭ 5) as well as by combined treatment with EPA and insulin (307 Ϯ 30 DU, p Ͻ 0.01, n ϭ 5) (Fig. 3A). No additive increase of GRB2 associated with IRS-1 was noticed when the cells were treated with both EPA and insulin (Fig. 3A). Similarly treated cells were analyzed for the association of PI3K with IRS-1. The amount of PI3K associated IRS-1 in vehicle-treated cells (78 Ϯ 11 DU) was increased either by EPA (376 Ϯ 29 DU, p Ͻ 0.01, n ϭ 5) or by insulin (530 Ϯ 45 DU, p Ͻ 0.01, n ϭ 5) as well as by combined treatment with EPA and insulin (662 Ϯ 47 DU, p Ͻ 0.01 versus basal; p Ͻ 0.01 versus insulin-treated cells, n ϭ 5) (Fig. 3B). An additive increase of PI3K associated with IRS-1 was noticed by combined treatment with EPA and insulin (Fig. 3B).
Furthermore, since MAPKs and Akt are downstream substrates for GRB2 and PI3K, respectively, we tested whether MAPKs and Akt are involved in cellular responses to EPA. Phosphorylated active MAPK was collected from cell lysates using anti-phospho-MAPK antibody, and its activity was determined by the amount of phosphorylated Elk1 fusion protein.
HepG2 cells were pretreated with 100 M EPA for 24 h, followed by stimulation with 100 nM insulin for 3 min, and analyzed for the MAPK activity. MAPK activity in vehicle treated cells (917 Ϯ 64 DU) was increased by insulin (1652 Ϯ 156 DU, p Ͻ 0.01) and conversely decreased by EPA (291 Ϯ 23 DU, p Ͻ 0.01) as well as by combined treatment with EPA and insulin (710 Ϯ 61 DU, p Ͻ 0.01 versus vehicle; p Ͻ 0.01 versus insulintreated cells, n ϭ 5) (Fig. 4A). These findings were incompatible with GRB2-associated IRS-1 but compatible with tyrosine phosphorylation of SHC. We also measured the Akt kinase activity determined by the amount of phosphorylated glycogen treated cells, n ϭ 5) (Fig. 4B). These findings were correlated with PI3K-associated IRS-1.
Next, we examined the mechanism by which EPA stimulates the tyrosine phosphorylation of IRS-1. EPA is a natural ligand for peroxisome proliferator-activated receptors (PPARs) and activates all three PPARs (PPAR-␣, PPAR-␦, and PPAR-␥) with the highest affinity to PPAR-␣. We tested the effect of other PPAR ligands on the tyrosine phosphorylation of IRS-1 in HepG2 cells. The effective dose of PPAR-␣-specific ligand (clofibrate), PPAR-␦ ligand (cPGI 2 ), or PPAR-␥-specific ligand (troglitazone) was chosen based on recent studies (20 -22). None of these ligands affected the amount of tyrosine-phosphorylated IRS-1 (Fig. 5A).
It has been reported that high intracellular glucosamine concentrations increase leptin production in adipose tissue (9). We also reported that EPA up-regulates leptin mRNA expression and its secretion through hexosamine biosynthetic pathway in 3T3-L1 adipocytes (8). Increased inflow of free fatty acid to HepG2 cells produces increased amounts of fatty acyl-CoA, causing the inhibition of glycolysis, and subsequently increases fluctose 6-phosphate, which increases the substrate for glutamine:fructose-6-phosphate amidotransferase, which plays a critical role as the rate-limiting enzyme in glucosamine biosynthesis. Glutamine:fructose-6-phosphate amidotransferase enhances the production of glucosamine 6-phosphate and its metabolites formed by subsequent acetylation and uridylation of glucosamine 6-phosphate (9,11,12). EPA would increase intracellular glucosamine 6-phosphate through this mechanism. DON, a well established inhibitor of glutamine:fructose-6-phosphate amidotransferase (23), however, did not affect EPA-induced tyrosine phosphorylation of IRS-1 (Fig. 5B).
To assess whether the step of protein synthesis is involved in stimulation by EPA of the tyrosine phosphorylation of IRS-1, HepG2 cells were treated with 100 M EPA in the absence or presence of 20 g/ml cycloheximide (CHX), a protein synthesis inhibitor, for 24 h, and IRS-1 was analyzed by immunoblot analysis with antibodies to phosphotyrosine. CHX blunted EPA-induced tyrosine phosphorylation of IRS-1 (Fig. 5C), although the amount of IRS-1 was decreased by CHX treatment.
We also investigated whether EPA affects glucose homeostasis in cell culture. Hepatic and renal gluconeogenesis is a major factor in maintaining glucose homeostasis. The rate-limiting enzyme of gluconeogenesis is PEPCK. This enzyme has no known allosteric control and is down-regulated by insulin at the transcriptional level. The rat hepatoma cell line H4-II-E cells have been used successfully to study the regulation of PEPCK expression, whereas HepG2 cells do not express PEPCK efficiently (24 -26). The amount of PEPCK mRNA in H4-II-E cells was reduced by EPA treatment and, in contrast, increased by arachidonic acid treatment as compared with control cells (Fig. 6). This observation is compatible with the result that EPA stimulated the IRS-1-PI3K pathway.
Next we tested the effects of EPA on cell proliferation, considering the difference between EPA and insulin with regard to the regulation of MAPK activity in HepG2 cells. We found that EPA inhibited proliferation of HepG2 cells in a time-and dosedependent manner (Fig. 7, A and B). The number of HepG2 cells was decreased by ϳ30% following a 72-h incubation with 100 M EPA (Fig. 7A). The phenotypic features were not changed. To exclude the possibility that EPA acted as a death factor and accelerated cell death of HepG2 cells, we counted the numbers of live and dead cells. The ratio of dead cells to the total cell number of EPA-treated cells was not significantly different from that in vehicle-treated cells (Table I). Furthermore, to investigate whether the decrease in the cell number was linked to apoptosis, we tested the DNA fragmentation showing the typical oligosomal "ladder" configuration by ApoLadder EX (Takara Biomedicals). There were no ladder formation in the cells (data not shown). EPA-inhibited cell proliferation was dose-dependent (Fig. 7B), and the result of the cell proliferation assay using MTS solution (Fig. 7C) was correlated well with that of the decrease in cell number induced by EPA. This observation is compatible with the finding that EPA inhibited MAPK activity in HepG2 cells. We also tested the effect of other fatty acids, docosahexaenoic acid, arachidonic acid, oleic acid, and palmitic acid on proliferation of HepG2 cells. Other fatty acids, except EPA, did not inhibit cell proliferation (Fig. 7D). Furthermore, we tested the effects of EPA on cell proliferation in other cell lines, rat L6 myocytes, mouse 3T3-L1 preadipocytes, and the human gastric adenocarcinoma cell line, MKN 45. EPA did not inhibit cell proliferation of these cells (Fig. 7E). DISCUSSION We demonstrated for the first time that EPA treatment for 24 h caused stimulation of both the basal and insulin-induced IRS-1-PI3K pathway in HepG2 cells as well as reduction of the amount of PEPCK mRNA expression in H4-II-E cells. Furthermore, it is of great interest that tyrosine phosphorylation of IRS-1 is specific for EPA, since other fatty acids cannot induce tyrosine phosphorylation of IRS-1. These findings are consistent with recent reports that EPA improves insulin sensitivity (1)(2)(3)(4). Moreover, it is our surprise that EPA inhibited MAPK activity as well as proliferation of HepG2 cells.
We focused on clarifying the mechanism by which EPA stimulated tyrosine phosphorylation of IRS-1. First, we examined the possibility of an involvement of PPARs. PPARs are nuclear receptors for fatty acids that regulate glucose and lipid homeostasis. The hypolipidemic actions of the fibrates appear to be primarily mediated through PPAR-␣, whereas the glucose-lowering effects of the thiazolidinediones are mediated through PPAR-␥ (27). EPA binds efficiently all three PPARs and activates these receptors in cell-based assays (20,22). However, neither PPAR ligands mimicked the tyrosine phosphorylation of IRS-1 by EPA, indicating that PPARs probably are not involved in this mechanism, although the involvement of PPAR-␦ is not completely ruled out. Human PPAR-␦-specific ligands have not been identified so far, and cPGI 2 , a ligand of PPAR-␦ and PPAR-␣, failed to stimulate tyrosine phosphorylation of IRS-1. Moreover, neither the function nor the array of genes regulated by PPAR-␦ is completely known. Therefore, the role of PPAR-␦ on the tyrosine phosphorylation of IRS-1 is not conclusive in our study. EPA has lipid-lowering effects and enhances insulin sensitivity in human and rodents, which is possibly mediated through PPAR-␣ and PPAR-␥, since these effects are similar to those of fibrates and thiazolidinediones. We speculate that EPA can improve insulin sensitivity and dyslipidemia possibly through its stimulation of the insulininduced tyrosine-phosphorylated IRS-1 as well as PPAR activation in accordance with dietary EPA intake. The second possibility is the involvement of hexosamine biosynthetic pathway, which mediates EPA-induced leptin expression as shown in our previous report (8). In this study, however, DON failed to affect EPA-induced tyrosine phosphorylation of IRS-1, suggesting that the involvement of hexosamine biosynthetic pathway in this mechanism is unlikely. Therefore, the precise mechanism of tyrosine-phosphorylation of IRS-1 by EPA remains unknown. The only positive result in this study is that CHX blunted EPA-induced tyrosine phosphorylation of IRS-1. We speculate that EPA could increase tyrosine phosphorylation of IRS-1 through endogenous protein synthesis or that the quantity of unknown EPA-stimulated kinase could be decreased, although the nature of this molecule remains to be elucidated.
IRS-1 is characterized to possess the 20 -22 potential tyrosine phosphorylation sites that are conserved between IRS-1 homologs. The surrounding amino acid residues are also highly conserved, and several of these represent potential binding sites for proteins that contain Src homology 2 domains (28,29). IRS-1 interacts with many Src homology 2 proteins with diverse phosphotyrosine motif requirements including PI3K and GRB2 (19,28). PI3K and GRB2 bind to different phosphotyrosine residues with IRS-1 (30,31). In the present study, EPA stimulated tyrosine phosphorylation of IRS-1, binding of PI3K to IRS-1, and Akt kinase activity and down-regulated the amount of PEPCK mRNA expression. These results are compatible with the actions of insulin-mimetics. Unlike insulin action, however, EPA inhibited both MAPK activity and proliferation of HepG2 cells irrespective of the presence of insulin, although it increased the association of GRB2 with IRS-1. The dissociation of the downstream molecules in the IRS-1-GRB2-MAPK pathway remains difficult to explain, but it may suggest the presence of a potent inhibiting mechanism by EPA of MAPK activity even under full activation by IRS-1-GRB2 association. It is possible that this is in part mediated by the inhibition by EPA of tyrosine phosphorylation of SHC, since phosphorylated SHC forms a complex with GRB2 and association of the SHC-GRB2 complex with the Ras guanine nucleotide exchange factor (Ras-GEF) mediates the localization of Ras-GEF to the plasma membrane (32). Once at the plasma membrane, Ras-GEF activates Ras by catalyzing the Ras-GTP for Ras-GDP exchange. This pathway may be inhibited through down-regulation of tyrosine phosphorylation of SHC by EPA. However, the mechanism by which EPA inhibited the tyrosine phosphorylation of SHC remains unknown. It is therefore likely that EPA affects tyrosine phosphorylation of SHC, MAPK activity, and cell proliferation by an as yet unknown pathway. Most important, the inhibition of cell proliferation by EPA is compatible with the result that EPA inhibited MAPK activity in HepG2 cells. Furthermore, other fatty acids, except EPA, did not inhibit proliferation of HepG2 cells, and EPA did not inhibit cell proliferation in other cell lines, L6 myocytes, 3T3-L1 preadipocytes, and MKN 45 cells. On the basis of these results, we propose that EPA may suppress cell proliferation, at least in HepG2 cells, by inhibiting the MAPK pathway.
Finally, the evidence that EPA caused not only up-regulation of IRS-1-PI3K pathway as well as down-regulation of gluconeogenesis but also inhibition of both the MAPK pathway and cell proliferation warrants further study to identify unknown mediators involved in EPA action cross-talking with the insulin signaling pathway.