Gnetin C from Melinjo helps restore metabolism by acting on fat tissue and liver in mice on a high-fat diet

Gnetin C, a resveratrol-like compound found in MSE and one of its key components, has previously been reported to exert beneficial effects on NAFLD²³. Therefore, in this study, we investigated whether Gnetin C could improve the pathological conditions of an HFD-induced obesity and diabetes model, similar to MSE. To induce obesity and diabetes, 4-week-old C57BL/6J mice were fed a high-fat diet (HFD) ad libitum for 10 weeks. Starting from week 6 of HFD feeding, Gnetin C (100 or 200 mg/kg) and MSE (1,000 mg/kg) were administered daily by oral gavage for 4 weeks (Fig. 1a). Body weight and fasting blood glucose levels were measured over time to evaluate the effects of Gnetin C and MSE. Both the Gnetin C- and MSE-treated groups showed reductions in body weight (Fig. 1b, c). Notably, Gnetin C at 100 mg/kg exerted weight loss comparable to MSE at 1,000 mg/kg, while Gnetin C at 200 mg/kg led to even greater weight reduction. In contrast, administration of Gnetin C at 200 mg/kg in wild-type mice did not produce the same stable weight reduction seen in the model mice—that is, no weight loss was observed under normal conditions—suggesting that the weight loss induced by Gnetin C is attributable to its alleviation of metabolic toxicity under high‐fat diet challenge (Supplementary Fig. 1). Similarly, fasting blood glucose levels were normalized in both Gnetin C- and MSE-treated groups (Fig. 1d, e). To assess the impact of Gnetin C and MSE on systemic oxidative stress induced by obesity and diabetes, levels of oxidative stress marker d-ROMs and antioxidant capacity (BAP) were measured in serum. Both Gnetin C and MSE significantly reduced oxidative stress and enhanced antioxidant capacity (Fig. 1f, g). These findings indicate that Gnetin C exerts anti-obesity and anti-diabetic effects similar to those of MSE, suggesting that Gnetin C is the key active component of MSE.

To evaluate the effects of Gnetin C and MSE on fat accumulation, subcutaneous and epididymal fat were excised at the time of dissection, and their weights were measured. Both Gnetin C- and MSE-treated groups exhibited reduced subcutaneous and epididymal fat mass (Fig. 2a-c). Although Gnetin C conferred metabolic improvements similar to MSE, it remains unclear whether their mechanisms of action are identical. To assess the effects of Gnetin C and MSE on circulating APN and high-molecular-weight (HMW) APN levels, ELISA was performed using mouse serum. Because APN is secreted from adipocytes, the ELISA data were normalized to fat mass. Both Gnetin C and MSE increased total APN levels, and notably, Gnetin C significantly elevated HMW APN levels (Fig. 2d, e); moreover, the extent of this increase was more pronounced than that of LMW APN (total APN minus HMW APN) (Fig. 2f), suggesting that APN multimerization plays a crucial role in the metabolic improvements conferred by Gnetin C.

Further analysis revealed that Gnetin C did not alter the expression of endoplasmic reticulum protein 44 (Erp44), another factor involved in APN multimerization²⁴, in adipose tissue. However, Gnetin C significantly increased the expression of DsbA-L and endoplasmic reticulum oxidoreductase 1α (Ero-1α)²⁵, both key factors that promote APN multimerization (Fig. 2g, Supplementary Fig. 2a, b). Additionally, Gnetin C enhanced the expression of peroxisome proliferator-activated receptor γ (PPARγ)²⁶, a known upstream regulator of DsbA-L and Ero-1α (Fig. 2h-j, Supplementary Fig. 3a, b). These findings suggest that Gnetin C promotes APN multimerization and exerts metabolic benefits by activating the PPARγ-DsbA-L-Ero-1α axis. Beyond these multimerization-related factors, we also assessed a panel of genes related to lipid metabolism. In adipose tissue, Gnetin C increased the expression of genes associated with fatty acid synthesis, adipocyte differentiation, lipolysis, and fatty acid β-oxidation (Fig. 2j).

To evaluate the effects of Gnetin C on hepatic lipid accumulation, H&E staining was performed. The results showed that Gnetin C reduced lipid accumulation in the liver (Fig. 3a, b). In addition, liver weight was not significantly affected by Gnetin C treatment (Fig. 3c). Next, to identify the key regulatory factors underlying the lipid-lowering effects of Gnetin C in the liver, we conducted a comprehensive analysis of genes involved in fatty acid synthesis, triglyceride synthesis, fatty acid β-oxidation, and oxidative stress. Among these genes, fibroblast growth factor 21 (Fgf21), which is associated with fatty acid β-oxidation, was significantly upregulated (Fig. 3d). FGF21, a hepatocyte secreted by the liver, is known for its potent metabolic regulatory effects on adipose tissue and other organs, making it a subject of intense research^27,28. Measurement of circulating FGF21 levels by ELISA, normalized to liver weight, revealed that Gnetin C further enhanced the HFD-induced increase in FGF21 secretion (Fig. 3e). Considering that obesity is associated with increased FGF21 secretion and FGF21 resistance²⁹, the additional upregulation of endogenous FGF21 may be crucial for the metabolic improvements observed in Gnetin C treatment.

To investigate the mechanism by which Gnetin C increases FGF21 expression, we focused on peroxisome proliferator-activated receptor α (PPARα)^27, a nuclear receptor known to regulate FGF21 expression. However, immunohistochemical analysis of PPARα in liver tissue showed that Gnetin C did not alter PPARα expression (Fig. 3f, g). We then turned our attention to sirtuin 1 (Sirt1), a NAD-dependent protein deacetylase known to induce FGF21 expression and improve fatty liver³⁰. Previous studies have shown that resveratrol, a Sirt1 activator, enhances AMP-activated protein kinase (AMPK) activation and mitochondrial function in a Sirt1-dependent manner³¹. Given that Gnetin C is a resveratrol dimer, we hypothesized that Gnetin C might similarly influence Sirt1 expression. Immunohistochemical analysis of Sirt1 in liver tissue revealed that Gnetin C increased Sirt1 expression (Fig. 3h, i), suggesting that Sirt1 upregulation may contribute to the increase in FGF21 expression observed with Gnetin C treatment.

It remains unclear whether the effects of Gnetin C on adipose and liver tissues are direct or mediated indirectly through APN or FGF21^32–34. To clarify this, we investigated the direct effects of Gnetin C in vitro. First, we used differentiated adipocytes derived from mouse 3T3-L1 preadipocyte. After 10 days of differentiation, 3T3-L1 adipocytes were treated with Gnetin C. Gnetin C not only reduced lipid accumulation (Fig. 4a, b), but it also produced a profile similar to the in vivo results: a slight increase in LMW APN levels in the culture supernatant, but a marked increase in HMW APN (Fig. 4c-e). These findings suggest that Gnetin C directly promotes APN multimerization in adipose tissue, thereby suppressing lipid accumulation.

Next, we used HepG2 cells, a human hepatoma-derived cell line treated with the saturated fatty acid palmitic acid to induce lipid accumulation. Similar to 3T3-L1 adipocytes, Gnetin C treatment reduced lipid accumulation in HepG2 cells (Fig. 4f and g). Moreover, the expression of Fgf21 and Sirt1 was upregulated (Fig. 4h-j, Supplementary Fig. 4a, b), suggesting that Gnetin C directly activates the Sirt1-FGF21 axis in liver tissue to suppress lipid accumulation.

Previous studies have reported that resveratrol enhances Sirt1 activity by directly binding to it³⁴. To investigate whether Gnetin C enhances Sirt1 activity through similar mechanisms, a Sirt1 activity assay was performed. This assay measures fluorescence emitted by acetylated substrates upon their deacetylation by Sirt1 in the presence of NAD⁺ and tissue or cell lysates (Fig. 5a). When liver lysates from Fig. 3 were tested, Sirt1 activity was found to be increased in the Gnetin C-treated group (Fig. 5b). Similarly, Sirt1 activity was elevated in lysates from HepG2 cells treated with Gnetin C (Fig. 5c), consistent with the increase in Sirt1 expression (Fig. 4i, j). These findings indicate that Gnetin C enhances Sirt1 activity in vivo, particularly in liver tissue. Next, to determine whether Gnetin C enhances Sirt1 activity through a direct binding mechanism, a Sirt1 activator screening assay was performed. This assay evaluates direct activation by incubating acetylated substrates and NAD⁺ with candidate direct activators in the presence of recombinant Sirt1 protein (Fig. 5d). As a positive control, the extent of Sirt1 activation by resveratrol was compared with a negative control (0.1% DMSO). Consistent with previous reports, resveratrol exhibited a concentration-dependent direct activation of Sirt1 (Fig. 5e). When tested under the same condition, Gnetin C also demonstrated a concentration-dependent direct activation of Sirt1, similar to resveratrol. Notably, under these conditions, Gnetin C induced a higher level of Sirt1 activation than resveratrol (Fig. 5f), indicating that Gnetin C can directly activate Sirt1 in vitro and may exhibit stronger activity than resveratrol in this context.

Previous studies have reported that FGF21 suppresses hepatic lipid accumulation through autocrine action³⁵, and it exerts paracrine effects on various organs. Among these paracrine effects, its impact on adipose tissue is particularly significant, as FGF21 has been shown to increase APN levels in adipose tissue, thereby improving glucose and lipid metabolism^36,37. Additionally, FGF21 has been reported to activate the PPARγ-DsbA-L-Ero-1α pathway³⁶ and increase HMW APN levels³⁸. These findings suggest that Gnetin C may exert its effects not only through direct action on adipose tissue but also by enhancing the paracrine effects of FGF21, promoting APN multimerization in adipose tissue. To investigate the potential role of FGF21-mediated autocrine and paracrine signaling in the metabolic improvements induced by Gnetin C, we analyzed the expression of FGF21 receptors in adipose and liver tissues. FGF21 receptors consist of a cell surface receptor complex comprising fibroblast growth factor receptor 1 (FGFR1) and the co-receptor protein βKlotho³⁹, and the expression levels of both components were examined. In liver tissue, Gnetin C increased FGFR1 expression but did not affect βKlotho expression (Fig. 6a, b). These findings, together with the results in Fig. 5, suggest that FGF21 upregulated by Gnetin C contributes to the suppression of hepatic lipid accumulation primarily through autocrine action. In adipose tissue, Gnetin C increased βKlotho expression but did not alter FGFR1 expression (Fig. 6c, d). This observation suggests that, in addition to Gnetin C's dual adipose and hepatic actions, FGF21 upregulated by Gnetin C may act on adipose tissue via paracrine signaling, activating the PPARγ-DsbA-L-Ero-1α axis and thereby partially contributing to APN multimerization (Fig. 6e).

MSE powder (YMP-M-181201), containing 0.07% RES, 2.41% gnetin C, 23.9% gnemonoside A, 4.51% gnemonoside D, and 9% dextrin as an excipient, was obtained from the Yamada Bee Company, Inc. (Okayama, Japan). Gnetin C (purity: >98%, HPLC) was prepared according to a previously described method¹¹.

C57BL/6J mice were housed in the animal experimentation facility at Kumamoto University in accordance with the guidelines of the university's Animal Facility Center. The mice were maintained under a 12-hour light/dark cycle at a room temperature of 20 °C. For HFD, we purchased the chow from Oriental Yeast Co., Ltd. (Tokyo, Japan), and replaced once weekly with 300 g of fresh chow after an overnight fasting period. The composition of the HFD was 25% casein, 20% safflower oil, 14% beef tallow, 8% dextrin, 7% lactose, 7% sucrose, and 5% cellulose. Five-week-old mice were randomly divided into five groups and fed with the HFD for six weeks. After ensuring that the average body weight of each group was similar at the end of the sixth week, the mice were fed HFD for an additional four weeks. MSE was administered once daily for four weeks at a dose of 1,000 mg/kg by oral gavage using a gastric tube. Similarly, Gnetin C was administered once daily for four weeks at doses of 100 mg/kg or 200 mg/kg by oral gavage using a gastric tube. The Gnetin C dose was determined based on the amount of Gnetin C and its glycosides present in 1,000 mg/kg of MSE. Notably, 1,000 mg/kg of MSE contains not only Gnetin C but also its glycosides (Gnemonosides), and assuming complete conversion of these glycosides to Gnetin C, while also accounting for inter-lot variations, the effective dose is estimated to fall within the range of 100 to 200 mg/kg. Therefore, two dose levels of 100 and 200 mg/kg were established. In the high-concentration group (200 mg/kg), gastrointestinal symptoms such as diarrhea and lymphangiectasia-like conditions were frequently observed in mice. Before administration, the test compounds were thoroughly suspended using a mortar and pestle. Blood glucose levels were measured using an Accu-Chek Compact glucometer (Roche, Basel, Switzerland), with blood samples collected via tail vein puncture. Fasting blood glucose levels were measured after more than 12 h of fasting, during which water was freely available. Anesthesia was induced in mice using an intraperitoneal injection of three types of mixed anesthesia (0.75 mg/kg of medetomidine [NIPPON ZENYAKU KOGYO CO., LTD., Fukushima, Japan], 4 mg/kg of midazolam [Sandoz K. K., Tokyo, Japan] and 5 mg/kg of butorphanol [Meiji Animal Health Co., Ltd., Kumamoto, Japan]). Following confirmation of a surgical plane of anesthesia, euthanasia was performed by exsanguination via cardiac puncture. These procedures adhered to the 2020 AVMA Guidelines for the Euthanasia of Animals. The animal study was conducted in accordance with the ARRIVE guidelines (Animal Research: Reporting of In Vivo Experiments). All animal procedures were approved by the Animal Welfare Committee of Kumamoto University (#A2024-050, to T.S.) and conducted in compliance with the relevant national and institutional guidelines for the care and use of laboratory animals.

3T3-L1 cells (RIKEN BRC, Japan) were used to evaluate the direct effects of Gnetin C on adipocytes. The cells were cultured in DMEM (High Glucose) medium (Wako, Osaka, Japan) supplemented with 10% Fetal Calf Serum (FCS) and 1% antibiotics (P/S; Penicillin G [100 U/mL] and Streptomycin [100 µg/mL]). To induce differentiation, the cells were maintained at confluence for two days, then the medium was replaced with 3T3-L1 adipocyte differentiation medium (KAC, Kyoto, Japan) and cultured for three days. Subsequently, the cells were cultured in a 3T3-L1 adipocyte culture medium (KAC, Kyoto, Japan) for seven days. HepG2 cells were obtained from the American Type Culture Collection (ATCC) and used to evaluate the direct effects of Gnetin C on hepatocytes. The cells were cultured in DMEM (Low Glucose) medium (Wako, Osaka, Japan) supplemented with 10% Fetal Bovine Serum (FBS) and 1% antibiotics (P/S; Penicillin G [100 U/mL] and Streptomycin [100 µg/mL]). To induce lipid accumulation in HepG2 cells, the cells were treated with 0.3 mM palmitic acid (Thermo Fisher, Massachusetts, USA) for 18 h. For maintenance culture, the culture dishes (3003 dishes, Falcon, Texas, USA) were pre-coated with Cell Matrix type I-C (Nitta Gelatin, Osaka, Japan) and incubated at 37 °C for at least 30 min before use. All cells were cultured in a humidified incubator at 37 °C with 5% CO₂.

Total RNA was extracted from mouse adipose and liver tissues, 3T3-L1 cells, and HepG2 cells using RNAiso Plus^®. For reverse transcription (RT) reactions, the PrimeScript^® RT Master Mix (Perfect Real Time) kit (Takara Bio, Shiga, Japan) was used according to the manufacturer's protocol. Subsequently, PCR reactions were performed using the resulting cDNA, TB Green Premix Ex Taq II (Tli RNaseH Plus) (Takara Bio, Shiga, Japan), and gene-specific primers under the following conditions: 95 °C for 3 min; followed by 40 cycles of [95 °C for 10 s, 65 °C for 1 min]; then a plate read. The reactions were carried out using a CFX Connect™ Real-Time PCR Detection System (Bio-Rad Laboratories, California, USA). To normalize mRNA expression levels across samples and wells, Gapdh was used as the internal control for mouse tissues, while 18 S rRNA was used for cells. The primer sequences for each target gene in mice are shown in Table 1.

Proteins were extracted from mouse adipose tissue and HepG2 cells using RIPA buffer (50 mM Tris-HCl [pH 7.5], 150 mM NaCl, 1 mg/mL sodium deoxycholate, 1 mM Na₃VO₄, and 1% protease inhibitor cocktail). Protein concentrations were measured using the BCA method. Next, proteins were separated on 8% or 10% SDS-PAGE and transferred to a PVDF membrane (250 mA, 1.5 h). The membrane was then blocked for 1 h with 0.05% PBS-Tween containing 5% skim milk. After washing in 0.05% PBS-Tween, the membrane was incubated at room temperature for 1 h with primary antibodies diluted in Can Get Signal^® Solution 1 (TOYOBO, Osaka, Japan). Following the primary antibody incubation, the membrane was washed with 0.05% PBS-Tween and incubated at room temperature for 1 h with secondary antibodies diluted in Can Get Signal^® Solution 2 (TOYOBO, Osaka, Japan). After this incubation, the membrane was washed with 0.05% PBS-Tween, treated with Super Signal^® West Pico chemiluminescent substrate (Thermo Fisher, Massachusetts, USA), and signals were detected using the Vilber Bio Imaging Fusion system (Vilber Lourmat, Collégien, France). The detected bands were quantified using ImageJ software. Primary antibodies used were anti-PPARγ (sc-7273, Santa Cruz, Texas, USA) and anti-Sirt1 (sc-15404, Santa Cruz, Texas, USA). Anti-Hsc70 (ADI-SPA-815, Enzo Life Sciences, New York, USA) and anti-vinculin (sc-73614, Santa Cruz, Texas, USA) were used as loading controls.

Serum levels of total APN and HMW-APN were quantified using a Mouse HMW & Total APN ELISA kit (ALPCO, New Hampshire, USA) according to the manufacturer's protocol. Similarly, FGF21 levels were measured using a Mouse/Rat FGF21 ELISA kit (R&D Systems, Minnesota, USA), following the manufacturer's instructions. For these measurements, serum was obtained by centrifuging blood collected from the inferior vena cava of mice at 5,000 rpm and 4 °C for 30 min. Cell culture supernatants were also used for the assays.

Plasma levels of diacron-Reactive Oxygen Metabolites (d-ROMs) and Biological Antioxidant Potential (BAP) were measured using the Free Carrio Duo system (WISMERLL, Tokyo, Japan) and the corresponding d-ROMs and BAP test kits (WISMERLL, Tokyo, Japan)⁵⁷, according to the manufacturer's protocol. For sample preparation, serum was obtained by centrifuging blood collected from the inferior vena cava of mice at 5,000 rpm and 4 °C for 30 min.

Mouse livers were excised, washed with PBS (-), and immersed in 10% neutral buffered formalin at room temperature for 12 h. Following formalin fixation, the tissues were washed three times with PBS (-) and dehydrated with ethanol. Subsequent steps, including paraffin embedding and H&E staining, were outsourced to K.I. Steiner (Kumamoto, Japan). Stained images were captured using a BZ-X710 microscope (Keyence, Osaka, Japan), and lipid droplet quantification was performed using ImageJ software.

After removing the culture medium from HepG2 and 3T3-L1 cells, the cells were fixed with 4% paraformaldehyde, then washed and dehydrated with 60% isopropanol. Subsequently, the cells were treated with a Oil red O staining solution and incubated in the dark for 20 min. After removing the staining solution, Milli-Q (MQ) water was added, and stained images were captured using a BZ-X710 microscope (Keyence, Osaka, Japan). ImageJ software was used for further analysis and quantification of the stained images.

Paraffin-embedded liver tissue sections prepared for IHC staining were deparaffinized and immersed in Target Retrieval Solution (DAKO, Glostrup, Denmark). Antigen retrieval was performed using an autoclave at 121 °C for 20 min. To prevent non-specific staining, the VECTOR M.O.M. Immunodetection kit (VEC, California, USA) was used. The sections were incubated overnight at 4 °C in the dark with a 1:100 dilution of an anti-PPARα antibody (sc-398394, Santa Cruz, Texas, USA) and an anti-Sirt1 antibody (sc-15404, Santa Cruz, Texas, USA). On the following day, the sections were treated with M.O.M. Biotinylated Anti-Mouse IgG Reagent (VEC, California, USA), and the reaction was visualized using a diaminobenzidine (DAB) substrate system (Nichirei, Tokyo, Japan). Stained images were acquired using a Nano Zoomer S20 Digital Slide Scanner (Nikon, Tokyo, Japan), and the number of PPARα⁺ and Sirt1⁺ cells was quantified using HALO software (Indica Labs, Corrales, USA).

SIRT1 activity was assessed using a fluorogenic peptide substrate bearing an N-terminal fluorophore and a C-terminal quencher. Upon deacetylation by active SIRT1 in the presence of NAD⁺, the concurrently supplied protease (Developer) cleaves the substrate, separating the quencher from the fluorophore and generating fluorescence, thereby enabling quantification of deacetylase activity. For the SIRT1 activity assay, MQ water and kit components (SIRT1 Assay Buffer, Fluoro-Substrate Peptide, NAD⁺, and Developer) were added to microtiter plate wells, followed by the addition of liver tissue and HepG2 cell lysates, and the plate was incubated at room temperature. Fluorescence was measured 60 min after lysate addition at Ex/Em = 350/460 nm using a BioTek Cytation 5 plate reader (Agilent Technologies, CA, USA). This protocol quantifies NAD⁺-dependent deacetylase activity present in lysates and thus reports SIRT1 activity.

For the SIRT1 activator screening assay, MQ water and kit reagents (SIRT1 Assay Buffer, Fluoro-Substrate Peptide, and NAD⁺) were dispensed into microtiter wells. Resveratrol (LKT Laboratories, MN, USA) and Gnetin C, prepared as previously described¹¹, were applied at 10 − 1,000 µM (each in 0.1% DMSO) or vehicle control (0.1% DMSO), after which Developer and recombinant SIRT1 were added, and the reaction was incubated at room temperature. Fluorescence was measured 10 min after enzyme addition at Ex/Em = 350/460 nm using a BioTek Cytation 5 plate reader. This protocol measures the direct effects of test compounds on recombinant SIRT1 deacetylase activity under defined, NAD⁺-dependent conditions; increases in fluorescence indicate activation, whereas decreases indicate inhibition, independent of cellular signaling contexts.

Experimental data are presented as the mean ± SEM. Statistical significance was determined using one-way ANOVA followed by Dunnett's post-hoc test, with a significance level set at p < 0.05. All statistical analyses were performed using GraphPad Prism 9 (GraphPad Software, California, USA).