The phosphorylated (p)-AMPK (cat. no. 2535) and AMPK (cat. no. 2532) antibodies were obtained from Cell Signaling Technology, Inc. PPAR-γ (cat. no. sc-7273), SREBP1 (cat. no. sc-13551) and β-actin (cat. no. sc-81178) antibodies were obtained from Santa Cruz Biotechnology, Inc.

The root of the herb Drynaria fortunei was purchased from Nanum Pharmaceutical Company (cat. no. HA1800100101). The herb (400 g) was extracted in 4 l hot water at 100°C for 4 h. The extract was freeze-dried and the yield was calculated at 17.5%; [dried extract weight (38.465 g)/dry starting material weight (219.8 g)] ×100 (%).

The present study followed the methods of Park et al (32). DRE powder was lyophilized, extracted with water at 100°C for 4 h and purified using filter papers under a vacuum rotary evaporator (EYELA-Tokyo Rikakikai Co., Ltd.); the residual powder was stored at −20°C until needed. The powder was used to generate a supplemented diet containing 10% DRE and 45% HFD (Research Diets, Inc.); 20 g DRE powder was mixed with 180 of HFD. Male C57BL/6 N mice (specific-pathogen-free grade; age, 8 weeks; weight, 20±2 g) were purchased from Dae Han Bio Link Co., Ltd. The mice were adapted to modified conditions for 1 week, and 30 healthy mice were then used in the present study. The mice were randomly distributed into the following three groups (n=10/group): Normal diet (CON), 45% HFD-induced (HFD), and 45% HFD-induced and 10% DRE-administered (HFD + DRE) groups. The mice were provided ad libitum access to food and water. Mice in the HFD + DRE group were provided a HFD for 4 weeks leading to HFD-induced obesity. Subsequently, the mice were fed a HFD supplemented with 10% DRE from week 5. Mice in the CON and HFD groups were fed normal diet and HFD, respectively for 9 weeks. The mice were maintained under a 12/12 h light/dark cycle, at a constant temperature of 22±2°C and relative humidity of 55±9%. Body weight and food intake were measured weekly. For sacrifice, mice were placed in a 9 l container; 100% of carbon dioxide was supplied to the container at a volume displacement rate of 30% per min (~3 l/min). The flow was continued until 1 min after breathing or heartbeat stopped. Subsequently, cervical dislocation was performed to ensure the animal was sacrificed. All procedures were conducted following the National Institutes of Health guidelines (33) and the present study was approved by the Ethical Committee for Animal Care and the Use of Laboratory Animals of Sangji University (approval no. 2019-11; Wonju, South Korea). At the end of the 10-week period, liver and adipose tissues were obtained, rinsed, weighed and stored at −80°C until further analysis.

After sacrifice, body weight gain was divided with food intake. And the value was expressed as percentage).

Relative epididymal white adipose tissues were calculated epididymal WAT weight divided to body weight.

Segments of liver or epididymal WAT (eWAT) were suspended in PRO-PREP™ protein extraction solution (Intron Biotechnology, Inc.) and incubated for 20 min at 4°C. Cell debris was removed via microcentrifugation (20,784 × g, 4°C for 30 min) followed by quick freezing of the supernatant. Protein concentration was determined using the Bio-Rad protein assay reagent (Bio-Rad Laboratories, Inc.) according to the manufacturer's instructions. Cellular proteins (30 µg) from homogenized adipose tissue were separated by 10–12% SDS-PAGE and electro-blotted onto a polyvinylidene fluoride membrane. The membrane was then incubated for 1 h with blocking solution (5% skim milk) at room temperature, followed by overnight incubation with primary antibodies (1:1,000) at 4°C. Blots were washed three times with 0.1% Tween 20/Tris-buffered saline (T/TBS) and were then incubated with horseradish peroxidase-conjugated secondary antibodies [Peroxidase AffiniPure Rabbit Anti-Mouse IgG (H+L); cat. no. 315-035-003; Peroxidase AffiniPure Goat Anti-Rabbit IgG (H+L); cat. no. 111-035-003; both from Jackson ImmunoResearch Laboratories Inc. 1:2,500 dilution] for 2 h at room temperature. Blots were again washed three times with T/TBS, and then developed via enhanced chemiluminescence (GE Healthcare). Densitometric analysis was performed using ImageJ ver 1.53a software (National Institutes of Health).

Total RNA was extracted from the liver tissue and eWAT using the EASY BLUE RNA extraction kit (iNtRON Biotechnology), according to the manufacturer's instructions. The RNA was then reverse transcribed into cDNA using an Maxime RT PreMix (Random primer, iNtRON Biochnology) according to the manufacturer's protocol. It was conducted using a GeneAmp PCR System 9700 (Applied Biosystems; Thermo Fisher Scientific, Inc.). The synthesized cDNA was 200 bp in size. A StepOnePlus Real-Time PCR System (Applied Biosystems; Thermo Fisher Scientific, Inc.) was used for amplification with Power-SYBR Green PCR Master Mix (Applied Biosystems). The qPCR thermocycling conditions were as follows: Preheating at 92°C for 2 min, followed by 50 cycles at 92°C for 30 sec, 60°C for 30 sec and 68°C for 30 sec. The expression data were calculated from the quantification cycle (Cq) value using the 2^−ΔΔCq method (34). GAPDH was used for normalization. The primer sequences used for RT-qPCR are listed in a previous study (35), as follows: AMPKα, forward (F) 5′-AGAGGGCCGCAATAAAAGAT-3′, reverse (R) 5′-TGTTGTACAGGCAGCTGAGG-3′; SREBP1, F 5′-ATCGCAAACAAGCTGACCTG-3′, R 5′-AGATCCAGGTTTGAGGTGGG-3′; PPARγ, F 5′-ATCGAGTGCCGAGTCTGTGG-3′; R 5′-GCAAGGCACTTCTGAAACCG-3′; GAPDH, F 5′-GACGGCCGCATCTTCTTGT-3′ and R 5′-CACACCGACCTTCACCATTTT-3′.

Briefly, 1.0–1.2 ml total blood was collected from each mouse using cardiac puncture following sacrifice. The collected blood was immediately centrifuged (1,000 × g for 30 min at 4°C) to obtain plasma. The plasma levels of triglycerides (TG) and total cholesterol (TC) were measured using commercial kits (AM157S-K for TG and AM 202-K for TC) (Asan Pharmaceutical, Co., Ltd.).

The liver tissue and eWAT from mice in each group were fixed in 10% buffered formalin for 5–10 min for eWAT as well as 20–30 min for liver at 37°C, embedded in paraffin and cut into 8-µm sections. Sections were stained for 6 h at 60–70°C with hematoxylin and eosin (H&E) for histological examination of lipid droplets. Adipocyte cell size was determined by measuring 10 randomly selected adipocytes per area of respective tissue and images were acquired using an Olympus SZX10 light stereomicroscope (Olympus Corporation).

Data are presented as the mean ± standard deviation of triplicate experiments. The experimental data were analyzed using one-way analysis of variance followed by Dunnett's post hoc test (GraphPad PRISM 5; Dotmatics). P<0.05 was considered to indicate a statistically significant difference.

Changes in body weight were tracked weekly for 10 weeks. The body weight of the HFD group was significantly increased after 4 weeks compared with that in the CON group (Fig. 1A). DRE was administered via the diet as 10% DRE supplemented in the HFD. The body weight of mice in the HFD + DRE group was lower than that in the HFD group. DRE induced a significant change in body weight at 6 weeks caused by adaptation of mice to the new diet. After 10 weeks, the body weight of mice in the HFD group was significantly increased (35.50±2.76 g) compared with that in the CON group (28.85±0.97 g). In addition, the HFD + DRE group exhibited reduced body weight (31.85±1.95 g) compared with that in the HFD group. Mice were allowed ad libitum access to the diet, and food intake was evaluated, showing no differences between the groups (Fig. 1B). To calculate the food efficiency ratio, the equation used to determine the protein efficiency ratio (PER; %) was adapted as follows: Body weight gain (g)/food intake (g) ×100 (36). Instead of evaluating the protein quality in food, the present study aimed to assess total food efficiency, which was significantly increased in the HFD group compared to the CON group as well as significantly decreased in the HFD + DRE group comparing to the HFD group (Fig. 1C). At the end of the experiment, after 10 weeks, anatomical examination confirmed the reduced body fat mass in the HFD + DRE group compared with that in the HFD group (Fig. 1D). In addition, after 9 weeks, the levels of serum biochemical markers, TG and TC, were significantly reduced in the HFD + DRE group compared with those in the HFD group and were significantly increased in the HFD group compared with those in the CON group (Fig. 1E and F).

Adipose tissue grows via two processes, hypertrophy and hyperplasia (37). In the present study, eWAT exhibited marked changes in size (Fig. 2A) and weight (Fig. 2B). The average weight of eWAT in the HFD group (1.90±0.32 g) was significantly increased compared with that in the CON group (0.70±0.09 g). This increase was suppressed by DRE (1.17±0.32 g) in the HFD + DRE group. This tendency was verified by determining relative eWAT weight (tissue weight/body weight) (Fig. 2C). To determine the effects of DRE on adipose tissue growth and lipid accumulation, H&E staining was performed on eWAT (Fig. 2D). As shown in Fig. 2D and E, the HFD group exhibited an increase in adipocyte size (113.22±4.57 µm) compared with that in the CON group (63.86±6.30 µm); however, mice in the HFD + DRE group exhibited decreased adipocyte diameter (81.93±3.61 µm) compared with that in the HFD group.

Western blotting was conducted to clarify the effect of DRE on the expression levels of adipogenic transcription factors in eWAT. The HFD + DRE group exhibited restored protein expression of p-AMPK-α compared with that in the HFD group, which was significantly decreased compared with the CON group (Fig. 3A and B). Given the regulatory effect of DRE on p-AMPK-α (Fig. 3A and B), the protein expression levels of adipogenic transcription factors, SREBP-1 and PPAR-γ, were determined. The increased protein expression levels induced by HFD were mitigated by DRE dietary treatment (Fig. 3C-E). To assess the mRNA expression levels of these factors, RT-qPCR analysis was performed. The mRNA expression levels were consistent with the protein expression levels. DRE upregulated AMPK activation, and a downregulatory effect on SREBP-1 or PPAR-γ expression (Fig. 3F).

Excessive fat accumulation in the liver leads to fatty liver diseases, such as nonalcoholic fatty liver disease (38). Therefore, the present study observed the size and color of liver tissue and compared its weight between groups. Bigger liver tissue of HFD group compared to CON group was presented and brighter liver tissue of HFD + DRE group comparing to CON or HFD group was shown (Fig. 4A), there was no statistically significant difference in liver weight among the three groups (Fig. 4B). However, histological examination with H&E confirmed that HFD-induced lipid accumulation was alleviated by DRE, resulting in smaller and less frequent lipid droplets, as indicated with black arrows (Fig. 4C).

The present study assessed the mRNA and protein expression levels of adipogenesis factors via RT-qPCR and western blot analysis, respectively. Unlike the changes in AMPK activation observed in eWAT and protein expression of liver, the difference in the mRNA expression levels of AMPK among the groups was not significant (Fig. 5A, B and F). Also, DRE treatment didn't significantly recover the protein expression of AMPK phosphorylation comparing to HFD group whereas apparently decreased phosphorylated AMPK expression in HFD group. However, the adipogenic transcription factor SREBP-1, and its direct target PPAR-γ, were markedly induced by HFD and suppressed by DRE dietary treatment (Fig. 5C-F). In Fig. 5B, D, and E, the relative density of protein expression was presented.