Effect and mechanism of ginsenoside Rg1-regulating hepatic steatosis in HepG2 cells induced by free fatty acid

ABSTRACT

Ginsenoside Rg1 (G-Rg1) is a bioactive phytochemical that has been found to be beneficial for the treatment of several diseases including nonalcoholic fatty liver disease (NAFLD). But there is a lack of literature reporting the effect of G-Rg1 on lipid metabolism balance in NAFLD. We investigated the effect and mechanism of G-Rg1 on lipid metabolism in vitro. We found that G-Rg1 decreased the levels of TG, TC, and MDA, and increased activity of SOD. Results of RT-PCR and western blotting showed that supplementation with G-Rg1 downregulated the expression of PPAR γ, FABP1, FATP2/5, CD36, SREBP1 c, and FASN, while the expression of PPAR ɑ, CPT1, ACOX1, MTTP, and ApoB100 was upregulated, after induction by a free fatty acid. Taken together, we conclude that G-Rg1 inhibits lipid synthesis and lipid uptake, and enhances lipid oxidation and lipid export to reduce hepatic steatosis of HepG2 cells by regulating PPAR ɑ and PPAR γ expression.

Graphical abstract

The pathway of G-Rg1 ameliorate lipid accumulation in hepatocytes in vitro.

KEYWORDS:

• Ginsenoside Rg1
• nonalcoholic fatty liver disease
• lipid metabolism
• hepatic steatosis

Nonalcoholic fatty liver disease (NAFLD) is a clinical syndrome characterized by excessive lipid accumulation in the liver and hepatic cellular degeneration, except for alcohol and other definite liver damaging factors [1]. The disease spectrum of NAFLD includes nonalcoholic fatty liver (NAFL) and nonalcoholic steatohepatitis (NASH), which can progress to cirrhosis and even liver cancer [2]. NAFLD has become one of the most common chronic liver diseases worldwide, which accounts for a quarter of the cases in the global population [3,4]. Currently, lifestyle interventions are still the main treatment strategy for NAFLD. Drugs such as insulin sensitizer (metformin, pioglitazone) and antioxidants (vitamin E) have been shown to have certain therapeutic effects on NAFLD, but their safety and side effects still need to be further evaluated [1,5]. Therefore, research and development for NAFLD therapeutic drugs are urgently needed.

As a central regulator of lipid homeostasis, the liver is responsible for orchestrating fatty acid synthesis, export, and subsequent redistribution to tissues [6]. These processes are closely regulated by a complex interaction between hormones, nuclear receptors, and transcription factors; if one or more of these pathways are destroyed, liver lipid homeostasis is disrupted [7]. Excessive hepatic lipid accumulation results from an imbalance between lipid acquisition and clearance, leading to increased lipid synthesis and uptake, and decreased lipid oxidation and export [8].

Increased de novo lipogenesis (DNL) is one of the obvious pathological features of NAFLD. Sterol regulatory element-binding proteins (SREBPs), a class of regulatory transcription factors stimulating DNL in the liver, can stimulate the expression of acetyl-coA carboxylase (ACCα) and the downstream fatty acid synthase (FASN), and thus regulate lipid metabolism [9]. Uptake of circulating fatty acid by the liver depends on their transmembrane transport, which involves a variety of proteins including the family of fatty acid transport proteins (FATPs) and scavenger receptor CD36. Fatty acids are intrinsically hydrophobic and therefore do not readily diffuse into the aqueous cytosol. Thus, intracellular binding proteins shuttle fatty acid between organelles and facilitate their uptake and metabolism [10]. During lipid oxidation in the liver, carnitine palmitoyltransferase (CPT1) and acyl-coenzyme A oxidase 1 (ACOX1), key rate-limiting enzymes, catalyze the oxidation of mitochondrial and peroxidase enzymes, respectively [11]. Apolipoprotein (ApoB100), a major component of very-low-density lipoprotein (VLDL) and low-density lipoprotein (LDL), is responsible for synthesis, assembly, and secretion of triglyceride (TG) rich VLDL and is essential for lipid transport; microsomal triglyceride transfer protein (MTTP) is a lipid transporter and major regulator of ApoB100 assembly and secretion in cells [12]. ApoB100 and MTTP play key roles in VLDL secretion and lipid homeostasis.

Ginsenoside Rg1 (G-Rg1) is one of the main active components in ginseng, which has anti-tumor, anti-inflammatory, anti-oxidation, anti-diabetes, and other pharmacological effects [13,14]. Because Most of the ginsenosides are poorly absorbed in the gastrointestinal tract, the concentration required for full pharmacological activity is difficult to achieve [15]. Oral ingested G-Rg1 does not undergo hydrolysis in the stomach and only a small portion is absorbed directly into the small intestine [16], its derivatives (ginsenoside Rh1, protopanaxatriol, or F1), generated by intestinal bacteria-mediated deglycosylation, are mainly responsible for its efficacy [16,17]. These derivatives, with better bioavailability and bioactivity, can be absorbed into the blood through passive diffusion and play a direct pharmacological role, and part of the derivatives can further play a role after fatty acid esterification in the liver [18,19]. Therefore, the deglycosylation process of G-Rg1 is crucial for its biological activity, and the efficacy of transformation and bioavailability of ginsenosides may be partly dependent on the intestinal microflora. However, owing to the diversity in resident microflora between individuals, the effects of orally ingested ginsenoside may differ [20].

Our previous study revealed that liver lipid deposition, endoplasmic reticulum stress (ERS), and inflammatory responses are significantly ameliorated by G-Rg1 in NAFLD model mice, suggesting that G-Rg1 has a positive effect on NAFLD treatment [21,22]. Therefore, the present study aimed to further explore the possible mechanism and target of G-Rg1 in ameliorating lipid metabolism in NAFLD on the basis of previous studies, so as to provide new ideas for NAFLD treatment.

Materials and methods

Cell culture

HepG2 cells (Procell, Wuhan, China) were cultured in Dulbecco’s Modified Eagle’s medium (DMEM) (Hyclone, Logan, UT, USA) containing 10% fetal bovine serum (FBS) (GIBCO, Grand Island, NY, USA) and 1% penicillin/streptomycin (Beyotime, Shanghai, China) at 37°C and 5% CO2. Logarithmic phase cells were seeded in 6-well plates at a density of 2.0 × 10⁶cells/well.

Free fatty acid (FFA)and/or G-Rg1 incubation

Palmitic acid (PA) (Sigma-Aldrich, St. Louis, MO, USA) was first dissolved in the solution of 1 M NaOH, and then mixed with 10% fatty-acid-free bovine serum albumin (BSA) (Solarbio, Beijing, China) at 70°C; Oleic acid (OA) (Sigma-Aldrich, St. Louis, MO, USA) dissolved in 10% BSA solution and then mixed with PA (PA: OA = 2:1) in DMEM to form FFA (1 mM). G-Rg1 (purity≥98%) (Meilunbio, Dalian, China) was dissolved in a solution of dimethyl sulfoxide (DMSO) (Beyotime, Shanghai, China) and diluted in DMEM. After the cell confluence reached 50%-60%, cells were treated with 10% BSA control or FFA (1 mM) in DMEM for 24 h and next treated with G-Rg1 (25/50 μM) for another 24 h.

Cytotoxicity assay

HepG2 cells were seeded in 96-well plates at 2 × 10⁴ cells each well. Post 24 h pre-treatment with FFA, the cells were further treated with G-Rg1 (25, 50, 75, 100, and 125 μM) diluted in the DMEM for another 24 h. Subsequently, cell viability was assessed using Cell Counting Kit-8 Assay (CCK8; Dojindo, Tokyo, Japan) Assay according to the manufacturer’s instructions. The cells were incubated with 10% CCK-8 solution for another 1 h at 37°C, and the absorbance of each well was measured at 450 nm using the Varioskan Flash Microplate Reader (ThermoFisherScientific, Waltham, MA, USA). Cell viability = [absorbance (experimental hole – blank hole)/(control hole – blank hole)] ×100%

Oil red O staining

HepG2 cells were fixed with 4% paraformaldehyde for 20 min at room temperature. Then, cells were rinsed in 60% isopropanol for 3 min and stained with freshly prepared Oil Red O staining solution (Oil Red O stock solution: deionized water = 3:2) for 20 min at room temperature. The cells were washed 3 times with double-distilled water and then observed and photographed under a microscope (Olympus, Tokyo, Japan). To quantify the Oil Red O contents, isopropanol was added to each group and the absorbance at 485 nm was measured using a microplate reader.

Biochemical measurements

Post FFA and G-Rg1 treatment, cells were broken using Soniprep150 ultrasonic crusher (SANYO, Osaka, JAPAN), and intracellular triglyceride (TG) of HepG2 cells was quantitatively tested by TG assay kit (BC0625, Solarbio, Beijing, China). Total cholesterol (TC) (BC1985), micro malondialdehyde (MDA) (BC0025), and superoxide dismutase (SOD) (BC0175) were determined using the corresponding assay kits (Solarbio, China). All of the experiments were performed following the manufacturer’s instructions.

Quantitative real-time polymerase chain reaction (RT-PCR)

After FFA and G-Rg1 treatment, total RNA was extracted by TRIzol reagent (ThermoFisher Scientific, Waltham, MA, USA). NanoDrop2000 spectrophotometer (ThermoFisher Scientific, USA) was used to detect the concentration and purity of total RNA, and the all-in-one cDNA Synthesis Super MIX (Bimake, Shanghai, China) was used for reverse transcription to cDNA. Reaction conditions: 42°C for 15 min, 85°C for 2 min, and 4°C for 5 min. 2× SYBR Green qPCR Master Mix (Bimake, China) was used for RNA determination on a fluorescence quantitative PCR instrument (Bio-Rad, Hercules, CA, USA). Reaction conditions: 95°C for 10 min; 95°C 15 s, 60°C 45 s, 40 cycles; 95°C for 15 s; draw the dissolution curve at 60°C for 60 s and 95°C for 15 s. GAPDH was used as an internal control and the relative expression levels of mRNA were calculated using the 2^−ΔΔCt method. The primer sequences were designed by Invitrogen (Carlsbad, CA, USA) and shown in Table 1.

Table 1. Primers for quantitative RT-PCR.

Gene	Forward (5ʹ-3ʹ)	Reverse (5ʹ-3ʹ)
GADPH	CTGGGCTACACTGAGCACC	AAGTGGTCGTTGAGGGCAATG
FABP1	GTGTCGGAAATCGTGCAGAAT	GACTTTCTCCCCTGTCATTGTC
FATP2	TACTCTTGCCTTGCGGACTAA	CCGAAGCAGTTCACCGATATAC
FATP5	CATGGCGTGACAGTGATCCT	CAGCCCGTAGTCCATTGCC
CD36	GGCTGTGACCGGAACTGTG	AGGTCTCCAACTGGCATTAGAA
SREBP1 c	GCCCCTGTAACGACCACTG	CAGCGAGTCTGCCTTGATG
FASN	AAGGACCTGTCTAGGTTTGATGC	TGGCTTCATAGGTGACTTCCA
ACCα	ATGTCTGGCTTGCACCTAGTA	CCCCAAAGCGAGTAACAAATTCT
CPT1	ATCAATCGGACTCTGGAAACGG	TCAGGGAGTAGCGCATGGT
ACOX1	AATCGGGACCCATAAGCCTTT	GGGAATACGATGGTTGTCCATTT
MTTP	TTCTCACAGTACCCGTTCTTGGT	TCCCCTGCCTGTAGATAGCCT
ApoB100	AAAGCCTTGCTGAAGAAAACCA	GAACCAGGAGGGCAGAAATGA
PPAR α	ATGGTGGACACGGAAAGCC	CGATGGATTGCGAAATCTCTTGG
PPAR γ	GGGATCAGCTCCGTGGATCT	TGCACTTTGGTACTCTTGAAGTT

Western blotting

HepG2 cells were harvested and resuspended in RIPA lysis buffer (Wanleibio, Shenyang, China) containing 1% protease inhibitor and 1% phosphatase inhibitors (Solarbio, China) for 30 min on ice, after which they were sonicated thrice (10 seconds each) with the ultrasonic crusher (SANYO, Japan). The cell lysates were centrifuged at 14,000 × g for 15 min at 4°C. The supernatants were collected and the protein concentration was determined by BCA Protein Quantification Kit (KeyGENBioTECH, Jiangsu, China). Then, it was diluted and balanced with 5× protein loading buffer and denatured by water bath at 99°C for 10 min. Equal amounts of proteins (20–30 µg/lane) from HepG2 cells were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (Bio-Rad, USA); and transferred to polyvinylidene difluoride (PVDF) membranes (ThermoFisher Scientific, USA). The membranes were blocked with blocking buffer (5% skim milk in Tris‑buffered saline (BOSTER, Wuhan, China) containing 0.5% Tween‑20 (PBST) (Bio-Rad)) for 1 h followed by incubation with primary antibodies (Peroxisome proliferators-activated receptor gamma (PPARγ)) (1663–1, 1:1000, 58kD, Proteintech), peroxisome proliferators-activated receptor ɑ (PPARɑ) (15,540–1, 1:800, 52–55kD, Proteintech), fatty acid translocase CD36 (CD36) (18,836–1, 1:1000, 75kD, Proteintech), fatty acid synthase (FASN) (10,624–2, 1:800, 280kD, Proteintech), carnitine palmitoyltransferase 1(CPT1) (15,184–1, 1:800, 86kD, Proteintech), peroxisomal acyl-coenzyme A oxidase 1 (ACOX1) (10,957–1, 1:800 ,50kD, Proteintech), apolipoprotein B100 (ApoB100) (20,578–1,1:800, 130–170kD, Proteintech),fatty acid binding protein 1 (FABP1) (ab7366, 1:500, 10–12kD, Abcam), fatty acid transport protein 2 (FATP2) (ab83763, 1:500, 70kD, Abcam), fatty acid transport protein 5 (FATP5) (ab97884, 1:500, 75kD, Abcam), microsomal triglyceride transfer protein (MTTP) (ab75316, 1:800, 100kD, Abcam); acetyl-coA carboxylase (ACCα) (3662S, 1:500, 280kD, CST), phospho-acetyl-coA carboxylase (p-ACCα) (3661S, 1:500, 280kD, CST), glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (WL01114, 1:1000, 36kD, Wanlei bio), sterol regulatory element binding proteins 1 c (SREBP1 c) (WL02093, 1:500, 68kD, Wanleibio) overnight at 4°C.The membranes were then washed with PBST three times for 10 min each and incubated with secondary antibodies (HRP-conjugated anti-rabbit IgG (A21020, 1:5000, Abbkine); HRP-conjugated anti-mouse IgG (A21010, 1:5000, Abbkine)) for 1 h at room temperature. After another three washes, the membranes were detected via enhanced chemiluminescence (ECL) (Millipore, Billerica, MA, USA) using ECL western blotting detection reagents (Bio-Rad, USA). The level of target protein band densities was quantified using the Image Lab 6.0 software. The western blotting analysis was performed in triplicate.

Data analysis

SPSS 20.0 software (IBM Corp. Armonk, NY, USA) was used for statistical analysis. Results were expressed as means ± standard deviation (SD). The difference between the two groups was analyzed by one-way ANOVA followed by the SNK-q test. P < 0.05 was considered statistically significant.

Results

Effect of G-Rg1 on HepG2 cell viability

The chemical structure of G-Rg1 is shown in Figure 1(a). G-Rg1 concentration had no significant effect on the viability of HepG2 cells at 25 μM, but the cell survival rate was only 88.7%±2.5% at 75 μM (Figure 1(b)). Therefore, G-Rg1 concentrations of 25 μM and 50 μM were selected for follow-up testing to avoid inhibition due to G-Rg1 cytotoxicity.

Figure 1. Effects of G-Rg1 on the viability of HepG2 cells. The structure of G-Rg1 (a). HepG2 cells were exposed to various concentrations of G-Rg1 (0, 25, 50, 75, 100,125 μM). After incubation for 24 h, cell viability was determined by CCK8 assay (b). Data represent mean ± SD (n = 3). *P < 0.05 (compared with the vehicle-treated control group).

G-Rg1 ameliorate FFA-induced lipid accumulation in HepG2 cells

Incubation with 1 mM FFA led to a large amount of lipid accumulation in HepG2 cells. Compared with the control group, the absorbance (485 nm) value and intracellular red lipid droplet aggregation, presented as large round particle, were significantly higher in the model group (Figure 2(a,b)). Compared with the model group, the absorbance (485 nm) at 25 μM and 50 μM G-Rg1 groups decreased to different degrees, and the intracellular red lipid droplets also decreased. The TG and TC contents were 4.71 and 1.18 times higher, respectively, in the model group than those in the control group (Figure 2(c,d)). The intracellular TG and TC contents in 25 μM and 50 μM G-Rg1 groups were significantly decreased, indicating that G-Rg1 could improve lipid deposition in liver cells.

Figure 2. Inhibitory effects of G-Rg1 on FFA-induced steatosis in HepG2 cells. The lipids accumulation was shown by Oil Red O staining (Original magnification ×200) and the absorbance was read under the microplate reader at 485 nm (a, b). Effects of G-Rg1 on cellular total cholesterol in FFA-induced HepG2 cells (c). The effects of G-Rg1 on cellular triglycerides in FFA-induced HepG2 cells (d). Values are expressed as mean ± SD (n = 3). ** P < 0.01, ***P < 0.001 (compared with the FFA-treated model group).

Effect of G-Rg1 on FFA-induced HepG2 cells MDA and SOD

We evaluated the antioxidant capacity of lipid by measuring SOD activity and determined MDA as a marker of lipid peroxidation. SOD content decreased and MDA content increased in the model group with that in the control group (Figure 3(a,b)). Compared with the model group, SOD content increased and MDA content decreased in the G-Rg1 group at different concentrations. The effect was most significant at 50 μM concentration, with an 87.6% increase in SOD activity and a 29.4% decrease in MDA content. Thus, G-Rg1 could enhance the lipid antioxidant capacity and weaken lipid peroxidation in HepG2 cells.

Figure 3. Effects of G-Rg1 on FFA-induced lipid peroxidation in HepG2 cells. Effects of G-Rg1 on MDA content (a) and SOD activity (b) in FFA-induced HepG2 cells. Values are expressed as mean ± SD (n = 3). **P < 0.01, ***P < 0.001 (compared with the FFA-treated model group).

Effect of G-Rg1 on expression of genes related to lipid synthesis in HepG2 cells induced by FFA

To further study the relationship between DNL and G-Rg1, western blot and RT-PCR were used to detect the expression of genes related to the SREBP1 c signaling pathway. SREBP1 c and FASN levels, detected by western blot, increased in the model group compared with those in the control group (Figure 4(a,c)), but decreased in 25 μM and 50 μM G-Rg1 groups compared with those in the model group. Additionally, there was no significant change in ACCα expression in the model group (Figure 4(b)), but the expression of both p-ACCα and p-ACCα to ACCα ratio was significantly decreased (Figure 4(b,c)). There was no significant change in ACCα expression after G-Rg1 treatment, but the expression of both p-ACCα and p-ACCα to ACCα ratio in the G-Rg1 group with the concentration of 50 μM was significantly decreased. RT-PCR results showed that the expression levels of SREBP1 c, FASN, and ACCα were increased in the model group compared with the control group (Figure 4(d)). Compared with the model group, the expression of SREBP1 c, FASN, and ACCα decreased in the Rg1 group with different concentrations, and the effect was most significant at the concentration of 50 μM. It was suggested that G-Rg1 could inhibit FFA-induced lipid synthesis in HepG2 cells.

Figure 4. Effects of G-Rg1 on DNL-related mRNA and protein expressions induced by FFA. Total proteins were extracted for western blot (a), quantified by bands intensity (b, c) and total RNA was isolated for RT-PCR analysis of SREBP1 c, FASN, ACCα (d). Data represent mean ± SD (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001 (compared with the FFA-treated model group).

Effect of G-Rg1 on expression of genes related to lipid uptake in HepG2 cells induced by FFA

Western blot results showed that the expression of CD36, FATP2, FATP5, and FABP1 in the model group were all increased compared with the control group (Figure 5(a)). Compared with the model group, the expression of CD36 and FATP5 was decreased in the 25 μM and 50 μM G-Rg1 groups (Figure 5(b)). The protein expression of FATP2 and FABP1 decreased in the 50 μM G-Rg1 group, but the decrease was not significant at 25 μM. Then, the expression of genes related to lipid uptake at the mRNA level was detected by RT-PCR (Figure 5(c)). Compared with the control group, the expression of CD36, FATP2, FATP5, and FABP1 were increased in the model group. After Rg1 treatment, CD36, FATP2, FATP5, and FABP1 mRNA expressions were all decreased. The change trend was consistent with the results of western blot, it was suggested that G-Rg1 could inhibit FFA-induced lipid uptake in HepG2 cells.

Figure 5. Effects of G-Rg1 on lipid uptake-related mRNA and protein expressions induced by FFA. Total proteins were extracted for western blot (a), quantified by bands intensity (b) and total RNA was isolated for RT-PCR analysis of CD36, FATP2, FATP5, FABP1 (c). Data represent mean ± SD (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001 (compared with the FFA-treated model group).

Effect of G-Rg1 on expression of genes related to lipid oxidation in HepG2 cells induced by FFA

We evaluated the effect of G-Rg1 on FFA-induced lipid oxidation in HepG2 cells by measuring the expression of CPT1 and ACOX1 at the protein and mRNA levels. CPT1 and ACOX1 expression, detected by western blot, was lower in the model group than in the control group (Figure 6(a)), and higher in the G-Rg1 group at different concentrations, with the most significant increase at 50 μM, compared with the model group (Figure 6(b)). The mRNA level, detected by RT-PCR, was consistent with the protein level (Figure 6(c)). CPT1 and ACOX1 mRNA expression were lower in the model group than in the control group, and higher in the 50 μM G-Rg1 group than in the model group. Thus, G-Rg1 could enhance the lipid oxidation of HepG2 cells induced by FFA.

Figure 6. Effects of G-Rg1 on lipid oxidation-related mRNA and protein expressions induced by FFA. Total proteins were extracted for western blot (a), quantified by bands intensity (b) and total RNA was isolated for RT-PCR analysis of CPT1, ACOX1 (c). Data represent mean ± SD (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001 (compared with the FFA-treated model group).

Effect of G-Rg1 on expression of genes related to lipid export in HepG2 cells induced by FFA

The results of western blot showed that compared with the control group, MTTP and ApoB100 protein expressions in the model group were decreased (Figure 7(a)). Compared with the model group, both MTTP and ApoB100 protein expressions were increased after treatment with G-Rg1 of concentration at 50 μM, while the increase of 25 μM concentration was not significant (Figure 7(b)). The results of RT-PCR showed that, compared with the control group, both MTTP and ApoB100 were decreased (Figure 7(c)). Compared with the model group, MTTP mRNA was increased in the G-Rg1 group at different concentrations, and ApoB100 mRNA was increased at 50 μM concentrations. The expression trends of MTTP and ApoB100 at translation and transcriptional levels are consistent, suggesting that G-Rg1 can enhance FFA-induced lipid export of HepG2 cells.

Figure 7. Effects of G-Rg1 on lipid export-related mRNA and protein expressions induced by FFA. Total proteins were extracted for western blot (a), quantified by bands intensity (b) and total RNA was isolated for RT-PCR analysis of MTTP, ApoB100 (c). Data represent mean ± SD (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001 (compared with the FFA-treated model group).

Effect of G-Rg1 on transcription factors PPARα and PPAR γ

PPARα protein and mRNA expression decreased in the model group compared with that in the control group, but increased in the 25 μM and 50 μM G-Rg1 groups compared with that in the model group (Figure 8(a-c)). Contrarily, PPAR γ protein and mRNA expression increased in the model group compared with that in the control group, but decreased in the 25 μM and 50 μM G-Rg1 groups compared with that in the model group. These results suggested that G-Rg1 could regulate the expression of PPARα and PPAR γ in HepG2 cells induced by FFA.

Figure 8. Effects of G-Rg1 on PPARα and PPARγ mRNA and protein expressions. Total proteins were extracted for western blot (a), quantified by bands intensity (b) and total RNA was isolated for RT-PCR analysis of PPARα, PPARγ (c). Data represent mean ± SD (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001 (compared with the FFA-treated model group).

Discussion

The pathogenesis of NAFLD has not been fully elucidated. Recent studies have shown that the occurrence and development of NAFLD are jointly affected by liver cell lipid accumulation, insulin resistance, oxidative stress, and other factors [8]. There have been reports that NAFLD begins with hepatic lipid accumulation, and significant hepatocyte lipid accumulation is a risk factor for disease progression. Hepatic lipid homeostasis is essential for the prevention and treatment of NAFLD [23]. However, previous studies have not reported the effect of G-Rg1 on lipid metabolism balance in lipid deposition cells. In the present study, lipid deposition in HepG2 cells was induced by 1 mM FFA, which is generally used for in vitro fatty liver cell model establishment [24,25]. However, a high concentration of FFA causes lipotoxicity, leading to apoptosis due to ERS, lysosomal dysfunction, or other factors. Moreover, lipotoxicity leads to β cell dysfunction in the pancreas and inhibits FFAR1expression in β cell, thus promoting insulin secretion [26]. Reportedly, 1 mM FFA induces a significant increase in lipid droplets in HepG2 cells without considerably affecting cell viability and apoptosis [27], and is thus the appropriate concentration for use with no significant lipotoxicity. We found that HepG2 cells treated with FFA showed a significant increase in TG and TC contents as well as lipogenesis. However, G-Rg1 treatment could reduce TG and TC content as well as lipid deposition in the cells, indicating that G-Rg1 could ameliorate FFA-induced HepG2 lipidosis.

Peroxisome proliferation-activated receptor (PPARs), with PPARα, PPAR β/δ, and PPAR γ, is a nuclear receptor involved in the transcriptional regulation of lipid metabolism, energy balance, and inflammation. PPARα is involved in lipid oxidation and lipid export, mainly expressed in the liver, brown adipose tissue, heart, and kidney; PPAR β/δ is commonly expressed in skeletal muscle, heart, gastrointestinal tract, and other tissues with high metabolism, which is mainly involved in the metabolism of fatty acid. PPAR γ is mainly expressed in adipose tissue and immune cells, and is responsible for regulating DNL and lipid uptake, and ameliorating insulin resistance [28].

PPAR γ expression is low under normal physiological conditions and gradually increases with the accumulation of lipids in the liver. It participates in the occurrence and development of NAFLD by regulating lipid synthesis and uptake [29]. Our study shows that G-Rg1 significantly decreased the high expression of DNL and lipid uptake-related genes, SREBP1 c, FASN, CD36, FATP2, FATP5, and FABP1, caused by FFA treatment. It is suggested that FFA could enhance the lipid uptake and synthesis in HepG2 cells, while the increase of fatty acid inflow would cause lipid overload in the cells. However, G-Rg1 could reduce DNL and lipid uptake in hepatocytes and reduce the lipogenesis of hepatocytes by decreasing the inflow of fatty acid. Previous studies have found that PPAR γ can promote the formation of NAFLD by regulating SREBP1 c to stimulate the expression of ACCα and FASN and upregulate the expression of CD36 and FABP1 to increase lipid uptake [30-32]. After the knockout of PPAR γ, the expression of lipid synthesis and uptake-related genes in hepatocytes was decreased, and the lipid accumulation of hepatocytes decreased as well [33]. We speculated that the role of G-Rg1 in ameliorating lipid uptake and synthesis in the NAFLD cell model may be related to the regulation of PPAR γ. Our study found that FFA treatment significantly increased the expression of PPAR γ in hepatocytes. Meanwhile, the expression of DNL and lipid uptake-related genes also changed with the change of PPAR γ expression, indicating that PPAR γ was an effective lipid-lowering target. G-Rg1 can reduce lipid uptake and DNL through the PPAR γ pathway, and it plays a role in hypolipidemic effects by reducing the inflow of lipids from the source, alleviating the treatment burden under the condition of lipid overload in liver cells.

PPARα is highly expressed in normal liver and decreased in NAFLD patients [34]. It can participate in the regulation of liver lipid metabolism by regulating the genes related to β-oxidation and the export of lipid [35]; its activation induces the expression of genes related to β-oxidation and export of lipid, which can decrease lipid deposition in the fatty liver [36,37]. Both ApoB100 and MTTP are key genes involved in the production of VLDL and secretion of TG, and play an important role in the lipid export from the liver [38]. PPARα, CPT1, and ACOX1 are associated with lipid accumulation in the liver; moreover, hepatocyte-specific peroxisome gene knockout or ACOX1 inactivation can also lead to oxidative stress [39,40]. Expression of PPARα, CPT1, and ACOX1 was also decreased. Our study found that FFA treatment increased the content of MDA and decreased the activity of SOD in HepG2 cells. MDA is a cytotoxic product of lipid peroxidation. SOD is an antioxidant, and its decreased activity disrupts the oxidant-antioxidant balance system and induces oxidative damage. Furthermore, G-Rg1 was found to ameliorate ERS in the liver of C57 mice fed a high-fat diet in our previous study [21]. FFA overload may destroy the balance of the oxidant-antioxidant system and ER homeostasis in hepatocytes. Excessive FFA accumulation would lead to a compensatory increase in β-oxidation in hepatocytes and the production of a large number of reactive oxygen species (ROS). The inability of hepatocytes to clear excess ROS, due to the insufficiency of intracellular antioxidants to compensate for the increased ROS, leads to an increase in oxygen-free radicals and peroxides, and thus oxidative stress [41]. This in turn may lead to the formation of unfolded or misfolded proteins, causing ERS, and eventually aggravating the lipid deposition and inflammatory reaction in hepatocytes. Therefore, although the liver cell apoptosis caused by 1 mM FFA is not apparent, it may cause cell damage, lipid-oxidation dysfunction, and promote lipid accumulation in hepatocytes. The deposition of TG promotes the production of O₂⁻ in the mitochondrial electron transport chain, and the accumulation of free radicals causes lipid peroxidation, leading to the imbalance in the oxidant-antioxidant system in the liver [42]. However, G-Rg1 could decrease MDA content and increase the SOD activity in liver cells, while simultaneously increasing the expression of PPARα, CPT1, and ACOX1 too. Thus, ERS was ameliorated and the oxidant-antioxidant system in liver cells was balanced. Additionally, the regulatory effect of PPARα on lipid decomposition is also related to lipid oxidation and export. PPARα has been proved to increase the secretion of ApoB100 by activating the expression of MTTP [35]. Our study found that G-Rg1 can upregulate the low expression of MTTP and ApoB100 caused by FFA treatment. This is consistent with the variation trend of PPARα and suggested that PPARα is another effective target of G-Rg1 for ameliorating lipid deposition in liver cells. Thus, G-Rg1 can enhance the β-oxidation and export of lipid through the PPARα pathway, improve the balance of lipid metabolism in cells, and enhance lipid processing to reduce lipid accumulation in liver cells.

However, the current data on lipid oxidation in NAFLD are inconsistent; yet, even in studies showing enhanced oxidation of lipid, augmented oxidation of lipid appears to be inadequate in clearing the liver of lipids [43]. After enhancing the activity of DNL, hepatic lipidosis and insulin resistance still appeared [44]. Mitochondrial and β-oxidative dysfunction have also been reported in NAFLD patients, and lipid accumulation in the form of TG is considered a protective and adaptive response to lipid overload [45]. Moreover, DNL and lipid oxidation also interact, and the increase of DNL will inhibit lipid oxidation. However, in the fatty liver, both fatty acid synthesis and β-oxidation can be simultaneously increased as a result of the subcellular distribution of ACCα activity [44]. These results suggest that when fatty acids are overstored in hepatocytes, the lipid-oxidation pathway is inhibited and the oxidation capacity is weakened. Further, the expression of PPARα and its regulated oxidation-related rate-limiting enzymes, CPT1 and ACOX1, were decreased. However, reduced lipid uptake and synthesis by hepatocytes may alleviate the burden on hepatocytes. The lipid-oxidation ability in hepatocytes was partially restored, and even increased compensatory, to speed up lipid treatment and reduce lipid accumulation. Therefore, G-Rg1 can enhance lipid oxidation and reduce lipid accumulation in liver cells through PPARα pathway. But this pathway may play a secondary role, and reduced lipid uptake and DNL of liver cells may also contribute to increased lipid β-oxidation.

It is worth noting that AMP-activated protein kinase (AMPK), as a central regulator of cell metabolism and energy balance, has been shown to upregulate the expression of PPARα while down-regulating the expression of PPAR γ, and regulate lipid metabolism by regulating the expression of lipid synthesis, oxidation, and export-related genes such as SREBP1 c, FASN, ACCα, CPT1, and ApoB100 [46-48]. Our previous studies also found that G-Rg1 ameliorated lipid deposition in NAFLD model mice by activating AMPK [22]. Combined with the findings of our study, we hypothesized that the effect of G-Rg1 on ameliorating lipid deposition and metabolism in liver cells through PPARα and PPAR γ activation might be related to the activation of AMPK, whether G-Rg1 regulates PPARs by activating AMPK remains to be investigated.

In conclusion, our study showed that G-Rg1 could ameliorate lipid accumulation in hepatocytes through at least two pathways in vitro: DNL and lipid uptake in liver cells can be reduced by decreasing the expression of PPAR γ, which decreased the inflow of lipid; while, increasing PPARα expression enhanced β-oxidation capacity and lipid export in liver cells, which increased the outflow of lipids. These two effects are complementary, and their positive effects are at least partly due to PPARα and its regulation of β-oxidation. The improvement of lipid uptake function may be the initiating factor in this series of reactions.

Author contribution

Substantial contributions to conception and design: Y Gao, WX Huang; Data acquisition, data analysis, and interpretation: Y Gao, YL Zhu, J Zhang; Drafting the article or critically revising it for important intellectual content: SJ Zhang, JJ Li, JQ Zhao, Q Xiao. All authors read and approved the manuscript.

Disclosure statement

No potential conflict of interest was reported by the authors.

Additional information

Funding

This work was supported by the Basic Research and Frontier Exploration Project of Chongqing Yuzhong District Science and Technology Bureau (NO.20190138)