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. 2026 Mar 16;59(7):364–371. doi: 10.5483/BMBRep.2025-0114

FOXO6 activation promotes hepatic steatosis via PPARα inhibition in aged male rats

Yeong Un Kim 1,#, Ji Hyun Park 1,#, Dae Hyun Kim 1,*
PMCID: PMC13424494  PMID: 41655988

Abstract

Aging contributes to hepatic steatosis by increasing de novo lipogenesis. The Forkhead box O6 (FOXO6) transcription factor links insulin signaling to lipid metabolism. Activated FOXO6 induces hyperlipidemia and decreases peroxisome proliferator-activated receptor alpha (PPARα), thereby promoting hepatic lipogenesis. In this paper, we describe the role of FOXO6 in hepatic steatosis in aged male rats and liver cells, and examine the relationship between FOXO6 and PPARα, and the functional consequences of their altered interaction. We find that FOXO6 induces lipid accumulation by inhibiting PPARα in aged male rat livers. Our data show that AKT signaling negatively regulates FOXO6-induced hepatic lipid accumulation, and that a key β-oxidation gene, PPARα, is decreased in aged livers. We further demonstrate that FOXO6 activation decreases PPARα expression and increases lipid accumulation. Furthermore, interaction between FOXO6 and PPARα promotes hepatic steatosis in aged males. Also, high glucose upregulates FOXO6, reduces β–oxidation gene expression, and increases cellular TG–mediated lipid accumulation. Transcriptional activation of FOXO6 by aging and high glucose causes lipid accumulation by downregulating PPARα and hyperglycemia-responsive genes in aged male rats and liver cell cultures. We provide evidence that age-related insulin resistance suppresses β–oxidation through interaction between FOXO6 and PPARα, thereby promoting hepatic lipid accumulation in aged male rats.

Keywords: Aged male, FOXO6, Interaction, Lipid accumulation, PPARα,

INTRODUCTION

Aging is characterized by the accumulation of physical, psychological, and social changes in an organism over time. The risk of developing diseases of aging, including diabetes and cancer, increases with the aging process (1). Older people are more prone to developing diabetes and obesity, which are strongly connected with insulin resistance (2). Insulin resistance is a pathological condition in which cells fail to respond to normal insulin signals that promote glucose uptake into tissues. As a result, hyperglycemia and hyperinsulinemia arise due to reduced insulin-stimulated glucose uptake and compensatory insulin secretion by beta-cells. In addition, diabetes is characterized by the increased production of glucose and triglycerides, contributing to high blood sugar and high triglycerides (3). The pathway linking the effects of hyperglycemia to insulin resistance and age-associated lipid accumulation has yet to be clearly defined. Therefore, the molecular mechanisms involving Forkhead transcription factor O6 (FOXO6) and peroxisome proliferator–activated receptor α (PPARα) in sex-specific aging require further exploration.

FOXO proteins, a subfamily of transcription factors that include FOXO1, FOXO3, FOXO4, and FOXO6, are known to regulate genes that control cellular metabolic pathways (4). AKT–mediated phosphorylation of FOXO, stimulated by growth factors or insulin, causes FOXO to move out of the nucleus, and into the cytoplasm (4, 5). FOXO is activated by two separate mechanisms: the reduction of growth factor signaling, which lifts AKT’s inhibitory effect, and high levels of fatty acids like palmitate, which utilize the c–Jun N–terminal kinase (JNK) pathway (6). JNK activates FOXO by phosphorylating IRS proteins, thereby restricting insulin–responsive PI3K/AKT signaling. However, the role of FOXO6 in hepatic lipid metabolism and its possible contribution to hypertriglyceridemia in aging males and females remain unclear. Here, the objective of this in vivo study was to determine the association between FOXO6 and PPARα-related molecules.

Peroxisome proliferator–activated receptors (PPARs), a family of nuclear receptors, are ligand–modulated transcription factors. PPARs heterodimerize with retinoid X receptors (RXRs) and bind to PPAR response elements (PPREs) in the promoter regions of specific target genes to regulate their transcription. Currently, three PPAR subtypes, PPARα, PPARβ, and PPARγ, have been identified. Among them, PPARγ, by promoting adipogenesis and managing lipid storage in adipocytes, also influences the expression of adipocyte-derived factors and adipocytokines, including leptin and adiponectin, thereby reducing lipotoxicity (7), and modulating Fatty liver disease (steatosis) (8). It was found that in db/db mice, FOXO6 elevated PPARγ expression in insulin-resistant livers, which, in turn, enhanced liver lipogenesis and led to fat accumulation (9). Growing evidence suggests that PPARα activation plays a role in controlling apoptosis, lipid metabolism, and inflammation within cardiovascular disease. Glucose-mediated apoptosis is prevented by the PPARα agonist, fenofibrate in vascular endothelial cells (VECs) (10). PPARα activation by agonists or PPARα overexpression regulates the apoptosis of vascular smooth muscle cells (VSMCs) (11). Further research is needed to understand the molecular relationship between FOXO6 and PPARα in the accumulation of lipids during aging, as this has not previously been reported. Dysregulation of lipid metabolism has been shown to be a factor in several metabolic diseases (12). PPARα directly controls the expression of genes for peroxisomal and mitochondrial fatty acid (FA) oxidation, influencing intracellular lipid levels, FA uptake, and triglyceride (TG) catabolism (13, 14). Studies have increasingly demonstrated the role of PPARα in metabolic syndrome, which includes conditions like diabetes, obesity, and dyslipidemia (14).

PPARα is predominantly found in tissues with high rates of fatty acid oxidation (FAO), including the various tissues (liver, heart, skeletal muscle, brown adipose tissue, kidney), though it is also a significant component in other cells and tissues, such as the intestine, vascular endothelium, smooth muscle, and immune cells (e.g., monocytes, macrophages, and lymphocytes) (13). PPARα coordinates metabolic shifts in response to feeding and fasting (15). PPARα regulates genes involved in peroxisomal and mitochondrial β–oxidation, FA transport, and hepatic glucose production (the latter being specific to rodents) (16). FA derivatives, produced through lipolysis, lipogenesis, or FA catabolism, serve as ligands for PPARα. Acyl–CoA oxidase 1 (ACOX1), the initial enzyme in the rate-limited peroxisomal β–oxidation pathway, has substrates that function as probable PPARα agonists. Evidence consistently shows that ACOX1 knockout in mice leads to increased peroxisome proliferation, the development of liver cancer, and up-regulation of PPARα target genes (17). The oxidized phospholipid fraction of oxidized low-density lipoprotein (LDL) serves as a ligand for PPARα, thereby enhancing its transcriptional activity and inducing the expression of its target genes, fatty acid transport protein 1 (FATP1), in human primary endothelial cells (18). When mice are fed a fat-depleted diet, liver-specific knockout of fatty acid synthase (FAS), an enzyme catalyzing the synthesis of FAs, results in hypoglycemia and liver steatosis; dietary fat or a synthetic PPAR agonist reversed the activation of PPARα by FAS-dependent de novo lipogenesis products (19).

In the present study, we examined age-related lipid accumulation in aged male and female rats, and assessed how FOXO6 and PPARα respond during hyperglycemia in hepatic tissues and cells. The intent of this is to define the molecular mechanisms underlying hyperlipidemia during aging in both sexes.

RESULTS

Changes in glucose, insulin, and lipid content in aged male and female rats

Obesity is generally accompanied by changes in circulating lipid levels due to chronic liver disease (19). Aging male and female rats were subjected to lipid profile test. Aging significantly increased fasting glucose in male rats (Fig. 1A). Aged males also showed hyperinsulinemia and hypertriglyceridemia, with increased insulin (Fig. 1B), plasma non-esterified fatty acids (NEFA; Fig. 1C), and TG (Fig. 1D), compared with young groups of the same age and sex. In contrast, no significant changes were observed in aged females. These findings suggest that dyslipidemia and hyperglycemia are induced during aging in males.

Fig. 1.

Fig. 1

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Aging-related serum changes in insulin resistance and lipogenesis. (A) Fasting blood glucose levels. (B) Fasting plasma insulin levels. (C) Free fatty acid and (D) triglyceride levels in the serum of aged male and female rats (n = 7 per group). Data are expressed as the mean ± standard deviation. #P < 0.05; ###P < 0.001 vs. 6-month-old rats.

Changes in FOXO6 and hepatic TG in the aging liver

To examine the impact of the aging process on lipid profiles in male and female rats, we analyzed hepatic lipid parameters. FOXO6 phosphorylation was reduced in the livers of aged males (Fig. 2A), whereas immunohistochemistry showed its expression to be increased (Fig. 2B). TG content in liver homogenates was assessed to substantiate age-related changes in hepatic lipids. An increase in hepatic TG was observed in aging male groups (Fig. 2C). Oil Red O staining was used to visualize and confirm the findings (Fig. 2D).

Fig. 2.

Fig. 2

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Aging-related increase in FOXO6-induced lipid accumulation. (A) Western blot analysis of p–FOXO6 and FOXO6 protein levels in the livers of aged male and female rats. (B) Immunohistochemical staining for FOXO6 in aging liver. Scale bar: 200 μm. (C) Hepatic triglyceride levels in aged rats. Data are expressed as the mean ± standard deviation. ###P < 0.001 vs. 6 months. (D) Oil Red O staining of aged livers to visualize lipid accumulation. Scale bar: 50 μm.

Expression of PPAR genes in aged male liver

PPARs modulate insulin resistance and inflammasome formation (20), both of which increase with age. We next explored FOXO6’s impact on lipogenesis in aging. PPARα regulates lipid metabolism by activating FA β–oxidation, and directly inhibits inflammatory responses (21). Transcripts of β–oxidation genes, such as PPARα, carnitine palmitoyltransferase 1A (Cpt1a), and peroxisomal acyl–coenzyme A oxidase 1 (Acox1), were decreased in aged males (Fig. 3A). However, comparison revealed that aged rats possessed higher mRNA expression of PPARγ and its downstream targets, such as fatty acid synthase (FASN) and stearoyl–CoA Desaturase (SCD) (Fig. 3B). Aged livers showed reduced nuclear PPARα protein levels compared to young, whereas nuclear PPARγ levels were increased (Fig. 3C). Also, expression of PPARα target genes involved in β–oxidation, including Cpt1a and Acox1, was decreased in aged animals (Fig. 3D).

Fig. 3.

Fig. 3

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FOXO6 regulates PPARs in aging liver. (A) Real-time PCR analysis of mRNA levels of PPARα, Cpt1α, and Acox1. Data are expressed as the mean ± standard deviation. *P < 0.05; **P < 0.01; ***P < 0.001 vs. 6 months. (B) Real-time PCR analysis of PPARγ, Fasn, and Acc. Data are expressed as the mean ± standard deviation. *P < 0.05; **P < 0.01; ***P < 0.001 vs. 6 months. (C) Western blot analysis of nuclear PPARs in aging liver. TFIIB was used as the nuclear loading control. (D) Western blot analysis of ACOX1 and CPT1α in cytosolic liver extracts (20 μg protein).

Changes in insulin signaling and interaction between PPARα and FOXO6 in aged liver

We investigated the interaction between FOXO6 and PPARα in aged male and female livers. Immunoprecipitation (IP) showed that this interaction was increased in the aging liver (Fig. 4A). The liver is the central regulator of fasting glucose via insulin signaling; therefore, we investigated this pathway in aged rats. The phosphorylation of IRS1 at Ser307, which is indicative of insulin resistance, showed a marked increase, while phosphorylated AKT (Ser473) decreased in aging males (Fig. 4B). This suggests that insulin signaling suppresses PPARα and FOXO6 crosstalk in the aging liver.

Fig. 4.

Fig. 4

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Interaction between FOXO6 and PPARα via insulin signaling in aging liver. (A) Co-immunoprecipitation showing physical interaction between FOXO6 and PPARα. (B) Expression of pSer–IRS1, pTyr–IRS1, IRS1, p–AKT, and AKT. β–actin was used as the loading control for cytosolic fractions.

Effects of glucose-mediated lipid deposition in hepatocytes

We investigated insulin pathway in hepatocytes, as glucose regulation in the liver is primarily mediated by insulin. Examining the interplay between elevated blood glucose and excessive lipid deposition, serum deprivation HepG2 cells were treated with 25 mM glucose. In this case, high-glucose (HG) treatment enhanced FOXO6 level and decreased p–FOXO6 levels (Fig. 5A). We further assessed glucose-induced lipid accumulation in HepG2 cells. A notable dependence on glucose increased intracellular TG levels (Fig. 5B). Real-time PCR revealed that Cpt1a and Acox1 expression was decreased under HG conditions (Fig. 5C). Concurrently, PPARα–associated genes also showed decreased expression (Fig. 5D). To assess glucose–induced PPARα activity, a PPRE/PPARα-luciferase assay was performed, revealing that glucose reduced PPARα activity (Fig. 5E). Immunoprecipitation with FOXO6 antibody or a non-specific IgG showed that HG conditions reduced the interaction between PPARα and FOXO6 (Fig. 5F). The results clearly indicate that high glucose promotes lipid accumulation by suppressing β–oxidation-related gene expression.

Fig. 5.

Fig. 5

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High glucose induces hyperlipidemia through the suppression of PPARα expression. (A) Western blot analysis of p–FOXO6 and FOXO6 in nuclear extracts after HepG2 cells were treated with glucose (25 mM) for 6 h. TFIIB served as the nuclear loading control. (B) Cellular triglyceride content after glucose treatment (25 mM, 24 h) measured via colorimetric assay. Data are expressed as the mean ± standard deviation. ###P < 0.001 vs. untreated cells. (C) Real-time PCR analysis of Cpt1a and Acox1 mRNA levels. Data are expressed as the mean ± standard deviation. #P < 0.05; ##P < 0.01 vs. untreated cells. (D) Western blot analysis of nuclear PPARα in glucose-treated cells. TFIIB was used as the nuclear loading control. (E) Effect of glucose on PPARα activity. HepG2 cells in 48–well plates were transfected with 0.5 μg pcDNA, full-length human PPARα, and a PPRE–luciferase construct. After 24 h, cells were treated with glucose (25 mM) for 8 h, then harvested to assess luciferase activity. Data are expressed as the mean ± standard deviation. ###P < 0.001 vs. pcDNA; ***P < 0.001 vs. PPARα. (F) Interaction of FOXO6 and PPARα in liver cells. Co-immunoprecipitation was performed using rabbit pre-immune IgG, anti-FOXO6, or anti-PPARα antibodies.

Increased FOXO6 activity leads to hepatocyte lipid

Heightened FOXO activity has been documented under reduced insulin conditions (4). We investigated the effect of elevated FOXO6 expression on PPARα level in HepG2 cells. We compared the effects of vehicle with the effects of an activated FOXO6 mutant (FOXO6−CA) at a multiplicity of infection of 100 on cells. FOXO6−CA treatment led to a marked decrease in PPARα expression (Fig. 6A). Cellular TG levels were elevated in FOXO6−CA-treated cells, compared with controls (Fig. 6B). Real-time PCR revealed decreased expression of Cpt1a and Acox1 in FOXO6−CA-treated cells (Fig. 6C). Thus, FOXO6 activation suppressed the expression of β–oxidation genes linked to PPARα. These findings suggest that FOXO6 activation strongly promotes lipid accumulation in liver cells. 

Fig. 6.

Fig. 6

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FOXO6 regulates hyperlipidemia through the suppression of PPARα expression in FOXO6 virus-treated cells. (A) Western blot analysis of PPARα expression in HepG2 cells treated with FOXO6 (100 MOI, 24 h). (B) Cellular triglyceride content measured via colorimetric assay. Data are expressed as the mean ± standard deviation. ###P < 0.001 vs. untreated cells. (C) Real-time PCR analysis of Cpt1a and Acox1 mRNA levels after FOXO6 treatment. Data are expressed as the mean ± standard deviation. #P < 0.05; ##P < 0.01 vs. untreated cells. (D) Proposed mechanism by which FOXO6 suppresses PPARα to promote lipid accumulation.

DISCUSSION

Aging is a complex, multifactorial process that is associated with various diseases, including chronic diseases, non-communicable diseases (NCDs), and metabolic diseases (1). The annual growth of the aging population has coincided with a rising incidence of diabetes and obesity (2). Hepatic fat accumulation is a significant risk factor for the development of metabolic dysfunction-associated steatotic liver disease (MASLD) due to the overconsumption of calories, primarily from a high-fat diet (22). MASLD is also linked to age-related metabolic dysfunction, characterized by impaired glucose metabolism, dyslipidemia, and chronic low-grade inflammation (23). Previous studies have shown that lipogenesis gene level increases with aging, partially driven by the increased expression of FOXO1 and PPARγ, transcription factors that lead to lipid accumulation (24). In the current research, the aged group showed significantly increased TG of blood and insulin levels compared with that in young male rats (Fig. 1). In contrast, no significant differences were observed in aged female rats, compared to their younger counterparts. Histological staining and aged male showed significantly increased hepatic TG levels, whereas aged females did not (Fig. 2). These sex differences in lipid accumulation are likely due to hormonal and reproductive differences.

By acting as a pivotal link in the insulin/IGF-1 pathway, FOXO transcription factors promote longevity. They achieve this by increasing resistance to oxidative stress and reducing the oxidative damage that contributes to accelerated aging (25, 26). The antioxidative effects of FOXO family members stem from their ability to bind to the promoters of antioxidant enzymes and increase their expression when insulin/IGF−1 signaling is inactive (27). Evidence from this study indicates the crucial role of FOXO in aging and its associated diseases. FOXO has been shown to lessen the harmful effects of faulty, aggregate-forming proteins, which are implicated in diseases such as Alzheimer’s and Huntington’s, signifying a role in neurodegeneration (28, 29). In our study, FOXO6 activation was observed in aged male rats (Fig. 2), contributing to elevated plasma glucose and Free fatty acid (FFA) concentrations, which likely suppressed hepatic lipid deposition (Fig. 1). Females tend to accumulate fat in specific regions owing to reproductive needs and estrogen-associated effects, whereas males typically have lower body fat due to androgenic influences on muscle mass.

FOXO6 inhibited expression of the transcription factor PPARα, further highlighting the essential role of PPARα in promoting hepatic β–oxidation (Fig. 3). Hyperglycemia triggered lipid accumulation via FOXO6 activation (Fig. 5A). Our data suggest that FOXO6 activation suppresses Cpt1 and Acox1 expression in liver cells (Fig. 5C), leading to increased TG accumulation. This indicates that FOXO6 enhances hepatic lipid accumulation by upregulating lipogenesis-related genes (Fig. 5B). Evidence also links PPARα to aging (30); reduced PPARα expression or activity has been reported in aged kidney, heart, and spleen tissues (31, 32). Studies using PPARα–deficient mice provide further support: these mice exhibit accelerated aging phenotypes, reduced longevity, and more age-associated lesions (33, 34). The significant association of fatty liver with insulin resistance in both humans (35) and animal models (36, 37) suggests that insulin resistance contributes to the pathogenesis of obesity-related fatty liver disease. Although FOXO and PPARα are individually implicated in hepatic steatosis, their interaction in the liver is not fully understood.

Our findings show that high-glucose conditions reduce FOXO6−PPARα interaction (Fig. 5F). FOXO6 also suppressed PPARα transcriptional activity, as shown by PPRE–luciferase assays, demonstrating that FOXO6 reduces PPARα interaction, as well as directly repressing its activity (Fig. 5E). This FOXO6–mediated reduction in PPARα activity appears to partially contribute to increased hepatic lipid accumulation. Our results further demonstrate that PPARα can regulate hepatic fat accumulation by modulating FOXO6 activity.

Hyperglycemia inhibits IRS/AKT signaling, leading to FOXO6 activation in aging males (Fig. 6D). In vivo evidence suggests that activated FOXO6 and PPARα interact to induce hepatic steatosis. Future studies may investigate the potential of targeting this pathway to prevent hepatic steatosis.

MATERIALS AND METHODS

Materials

All chemical reagents were obtained from Sigma (St. Louis, MO, USA), unless otherwise stated. Western blot detection reagents were obtained from Amersham (UK). RNAzolTM B was obtained from TEL−TEST, Inc. (Friendswood, TX, USA). Antibodies against PPARγ, β–actin, TFIIB, p–AKT, total AKT, pS−IRS, pT−IRS, and IRS were obtained from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Antibodies against PPARα, CPT1α, and ACOX1 were purchased from Abcam (Cambridge, UK). Antibodies against FOXO6 and p–FOXO6 (Ser184) were obtained from Dr. Dong (University of Pittsburgh, Pittsburgh, PA, USA). Horseradish peroxidase-conjugated anti-rabbit and anti-mouse IgG antibodies were obtained from Amersham (UK). Horseradish peroxidase-conjugated anti-sheep/goat IgG antibodies were purchased from Serotec (Oxford, UK). Polyvinylidene difluoride (PVDF) membranes were obtained from Millipore Corporation (Billerica, MA, USA).

Animal experimental procedures

Young (6 months old) and aged (24 months old) male and female Sprague–Dawley rats were purchased from Samtako (Gyeonggi-do, Korea), and acclimated to the animal care facility for 7 days before the experiments. The animals were housed in an air-conditioned atmosphere under a 12-h light/dark cycle, and provided free access to standard rodent chow (Samtako) and water. The animal study was approved by the Institutional Animal Care Committee of Pusan National University, and was performed in accordance with the guidelines for animal experiments issued by that university (approval number PNU−2020−2583).

Cells

Human hepatocellular carcinoma HepG2 cells were obtained from the American Type Culture Collection (Rockville, MD, USA). HepG2 cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM) (Nissui Co., Tokyo, Japan) supplemented with 10% heat-inactivated (56°C for 30 min) fetal bovine serum (Gibco, Grand Island, NY, USA), 233.6 mg/ml glutamine, 72 μg/ml penicillin–streptomycin, and 0.25 μg/ml amphotericin B, with the pH adjusted to (7.4 to 7.6) with NaHCO3. Cells were maintained at 37°C in a humidified atmosphere containing 5% CO2. The medium was then replaced with fresh medium.

Biochemical analysis

Blood samples were collected from the rats in each group. Different kits were used according to the manufacturers’ instructions to determine the concentrations of the following metabolites in the serum: insulin (Shibayagi, Japan), glucose, non-esterified fatty acids (NEFA), and TG (Shinyang, South Korea).

Western blot analysis

Homogenized samples were boiled for 5 min in a gel-loading buffer (0.125 M Tris–HCl, pH 6.8, 4% sodium dodecyl sulfate (SDS), 10% 2–mercaptoethanol, and 0.2% bromophenol blue) in a 1:1 ratio. Equal amounts of total protein from each sample were separated by SDS–polyacrylamide gel electrophoresis (SDS−PAGE) on 10% acrylamide gels, and transferred to PVDF membranes at 80 V for 1.5 h in a semi-dry transfer system. The membranes were immediately placed in blocking buffer (5% nonfat milk in 10 mM Tris at pH 7.5, 100 mM NaCl, and 0.1% Tween 20). The blots were blocked at room temperature (RT) for 30 min. The membrane was incubated with a specific primary antibody at 4°C overnight, followed by incubation with a horseradish peroxidase-conjugated anti-rabbit antibody at RT for 1.5 h. Labeled antibodies were detected using WESTSAVETM (AbFrontier, South Korea). Pre-stained protein markers were used for molecular weight determination.

Hepatic lipid content

Liver tissues (20 mg) and cells were homogenized in 400 μl of HPLC-grade acetone. After an overnight incubation with agitation at RT, 50 μl aliquots of acetone–extracted lipid suspensions were used to determine triglyceride concentrations via the Infinity triglyceride reagent (Thermo Electron). Hepatic lipid content was defined as milligrams of triglycerides per gram of total liver protein, as earlier described (38).

RNA isolation and real-time quantitative reverse transcriptase PCR (qRT−PCR)

RNA was isolated from liver tissue using the RNeasy Mini Kit (QIAGEN, Valencia, CA, USA). qRT−PCR was performed to quantify mRNA levels using SYBR Green and a CFX Connect System (Bio-Rad Laboratories, Hercules, CA, USA). The following primers were used for amplification: R−PPARα: F, 5’-AGGAAGCCATTCTGCGACAT-3’ and R, 5’-CTCTGCAGGTGGAGCTTGAG-3’; R−ACOX1: F, 5’-TCAGCAGGAGAAATGGATGC-3’ and R, 5’-TGGAAGTTTTCCCAAGTCCC-3’; R−CPT1: F, 5’-AAGCTGTGGCCTTCCAGTTC-3’ and R, 5’-GGATGAAATCACACCCACCA-3’; R−PPARγ: F, 5’-CCAGGTGACCCTCCTCAAGT-3’ and R, 5’-CTGCAGCAGGTTGTCTTGGA-3’; R−ACC: F, 5’-GGCACTCTGATCTGGTCACG-3’ and R, 5’-GCTCCGCACAGATTCTTCAA-3’; R−FASN: F, 5’-AGTGAGTGTACGGGAGGGCT-3’ and R, 5’-GCTGGGACACATGTGATGGT-3’; H−ACOX1: F, 5’-AGGTCACAGCTGTCCAACCA-3’ and R, 5’-TTACCCAGCCCTGGCTTAAT-3’; H−CPT1: F, 5’-GGTCCAGGTAGAGCTCAGGC-3’ and R, 5’-GTGCTCTGAGGCCTTTGTCA-3’. All primers were purchased from IDT (Coralville, IA, USA).

Immunoprecipitation of tissue extracts

Liver tissue and cells were lysed in buffer containing 40 mM Tris–HCl (pH 7.6), 120 mM NaCl, 20 mM glycerophosphate, 20 mM NaF, 2 mM sodium orthovanadate, 5 mM ethylenediaminetetraacetic acid (EDTA), 1 mM phenylmethylsulfonyl fluoride (PMSF), 0.1% NP−40, leupeptin (2 μg/ml), aprotinin (1 μg/ml), and pepstatin A (1 μg/ml). Cell extracts were centrifuged at 12,000 × g at 4°C for 15 min, incubated overnight at 4°C with the appropriate antibody, and then with 50% protein A agarose slurry overnight at 4°C. Immunoprecipitates were washed three times with IP buffer, and proteins were analyzed by SDS−PAGE and Western blotting.

Histological analysis

Oil Red O staining was performed as previously described (39), using frozen tissues under optimal cutting temperature to visualize lipid accumulation.

Transfection and luciferase assay

PPARα activity was measured using the PPARα–Luc vector. Transfection was performed with Lipofectamine 2000 (Invitrogen). Briefly, 1 × 104 cells/well were seeded in 48-well plates. At ∼40% confluence, cells were treated with 1 μg DNA/0.5 μl Lipofectamine 2000 complexes in 500 μl medium containing 5% serum for 24 h, followed by infection with FOXO6−CA (200 MOI). After 24 h incubation, cells were cultured for an additional 24 h. Luciferase activity was measured using the Steady-Glo Luciferase Assay System (Promega, Madison, WI, USA) and a luminometer (GENious, TECAN, Salzburg, Austria).

Immunohistochemical analysis

Liver sections were treated with 0.6% H2O2 in Tris-buffered saline (TBS; pH 7.5) to block endogenous peroxidase for 15 min at RT. Sections were then incubated with TBS containing 0.1% Triton X−100 and 3% goat serum (TBS−TS) at 37°C for 1 h, followed by incubation with primary anti-FOXO6 antibody (courtesy of Dr. Dong) in TBS−TS overnight at 4°C. Sections were incubated with goat anti-rabbit IgG–HRP secondary antibody (Santa Cruz) at RT for 3 h, stained with DAB, mounted with Dako mounting medium (Dako, Glostrup, Denmark), and cover-slipped. Images were acquired using an Olympus IX71 microscope (Olympus, Tokyo, Japan). Livers were fixed in 10% neutral buffered formalin.

Statistical analyses

Comparisons among three or more groups were performed using one-way ANOVA, followed by Bonferroni’s post hoc test. A P–value < 0.05 was considered significant. Data are expressed as the mean ± SD. Analyses were conducted using GraphPad Prism 5 (GraphPad Software, San Diego, CA, USA).

ACKNOWLEDGEMENTS

This study was supported by a National Research Foundation of Korea (NRF) grant, funded by the Korea government (MSIT) (grant number: RS−2024−00339004). We thank the Aging Tissue Bank (Busan, Korea) for providing useful information.

Footnotes

CONFLICTS OF INTEREST

The authors have no conflicting interests.

AUTHOR CONTRIBUTIONS

DHY designed and performed the experiments, and assembled and wrote the manuscript; YUK and JHP supervised the experiments and reviewed the manuscript. All authors contributed to the interpretation of the results. All authors revised and approved the manuscript.

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