Neoshirakia japonica (Siebold & Zucc.) Esser [Euphorbiaceae] fruit extract was purchased from the Natural Product Central Bank (KPM025-071). The source fruits were originally collected from Ongnyong-myeon, Gwangyang-si, Jeollanam-do, Korea, in 2004 and then dried in the shade and concentrated, and a voucher specimen (KRIB 0003259) is kept in the herbarium of the Korea Research Institute of Bioscience and Biotechnology. 19 g of fruit powder was added to 1 L of 99.9% methyl alcohol (HPLC grade) and extracted using 30 ultrasonication cycles at room temperature in an ultrasonic extractor (SDN-900H, SD-Ultrasonic Co., Ltd., Korea), with 15 min of ultrasonication at 40 KHz and 1500 W followed by a 120 min standing period per cycle. Subsequently, the extract was filtered using a qualitative filter (No. 100, Hyundai Micro Co., Ltd., Seoul, Korea). The filtered extract underwent a first drying process in a water jacket-type dry oven (SCE&G, Kyung-gi, Korea) at 40 °C for 48 h. The dried sample was then subjected to a second drying step using a centrifugal vacuum concentrator (HANIL SCIENTIFIC INC. Seoul, Korea) at room temperature and 1,500 rpm under reduced pressure. To ensure complete drying, this step was extended up to 24 h as needed. For the third drying step, 300 μL of distilled water was added to each of the dried samples, followed by freezing in a deep freezer. The frozen samples were then lyophilized using a freeze-dryer (Ilsin BioBase, Korea) for up to 36 h, resulting in the final yield of 2.03 g of NJFE. The dried extract was immediately dissolved in dimethyl sulfoxide (DMSO; Sigma, Saint Louis, United States) at a concentration of 50 mg/mL and subsequently diluted to 2 mg/mL for experimental use. The 2 mg/mL solution was aliquoted into 30 µL portions and stored at −80 °C. Once thawed, it was neither re-frozen nor returned to −80 °C storage. For animal experiments, the dried extract was freshly dissolved in PBS immediately prior to use at a volume sufficient for 6 days of treatment (three injections, administered every other day). The prepared solution was stored at 4 °C and used as needed over a 6-day period. Any remaining volume after 6 days was discarded.

3T3-L1 preadipocytes were purchased from ATCC (Manassas, United States). The cells were cultured in a growth medium consisting of 10% new bovine calf serum and 1% (v/v) penicillin-streptomycin (Gibco, Paisley, United Kingdom) in Dulbecco’s modified Eagle’s medium with high glucose (DMEM-H) (Gibco). When cells reached approximately 80% confluency, they were either passed using trypsin-EDTA (TE) (Gibco) or seeded into six-well plates.

When 3T3-L1 preadipocytes reached 100% confluency, they were maintained for an additional 2 days to arrest further growth. Then, the medium was replaced with 10% fetal bovine serum (FBS) and 1% (v/v) penicillin-streptomycin in DMEM-H supplemented with methylisobutylxanthine (0.5 mM), dexamethasone (0.25 mM), insulin (1 μg/mL), and indomethacin (0.125 nM) (MDI; Sigma), and the cells were cultured for 2 days. After this, the medium was replaced with 10% FBS, 1% (v/v) penicillin-streptomycin, and insulin (1 μg/mL) in DMEM-H to induce lipid accumulation. The cells were cultured in this medium for 6 days, with the medium changed every 2 days. Thus, the mature adipocyte differentiation induction process lasted for a total of 8 days.

RAW264.7 macrophages were purchased from Korean Cell Line Bank (KCLB, Seoul, Korea). The cells were cultured in DMEM-H supplemented with 10% Fetal bovine serum (FBS) (Gibco) and 1% (v/v) penicillin-streptomycin. When the cells reached 70%–80% confluency in 100-mm culture dishes, they were seeded into six-well plates at a density of 1 × 10⁶ cells/well. After 24 h, the cells were treated with lipopolysaccharide (LPS; Sigma) at a concentration of 1 μg/mL to induce an inflammatory response.

To evaluate the cytotoxicity of the NJFE, adipocytes were seeded in a 96-well plate at a density of 3 × 10⁴ cells/well and maintained for 24 h. The following day, when the cells reached 80% confluence, they were treated for 48 h with NJFE at concentrations ranging from 0.25 to 32 μg/mL or for the control group, the same volume of DMSO as used for the 32 μg/mL NJFE group. After treatment, 50 μL of cell counting kit-8 (CCK-8) solution (Dojindo Molecular Technologies, Kumamoto, Japan) was added to each well, and the plate was incubated at 37 °C for 1–2 h. The absorbance at 450 nm was measured using a microplate reader (Tecan, Männedorf, Switzerland).

At the end of differentiation, adipocytes were washed twice with PBS and fixed with 4% paraformaldehyde (Biosesang Inc., Yongin-si, Korea) for 1 h in the dark. After fixation, the cells were washed once with PBS. An oil red O (ORO) solution was prepared by diluting ORO (Sigma) with triple-distilled water at a 2:3 ratio, followed by centrifugation at 4,000 rpm for 20 min to collect the supernatant. The ORO solution was then added to each adipocyte sample, and the mixture was incubated for 30 min in the dark to allow staining. After staining, the cells were washed three times with triple-distilled water to remove any residual stain and then imaged under a microscope. After imaging, any remaining distilled water was removed by drying. To quantify the ORO content, 1 mL of isopropyl alcohol was added to each well to dissolve the stain, and the absorbance was measured at 450 nm using a microplate reader.

Total RNA was extracted from adipocytes and adipose tissue using RNAiso Plus (TRIzol; TaKaRa Bio, Kyoto, Japan), and the extracted RNA was converted into complementary DNA (cDNA) using a ReverTra Ace™ qPCR RT Master Mix kit (Toyobo, Kyoto, Japan). For the real-time reverse-transcription polymerase chain reaction (RT-qPCR), 1,000 ng of cDNA was mixed with SYBR Green master mix (Toyobo) and gene-specific primers, custom-designed and synthesized by Macrogen (Seoul, Korea) (Supplementary Table S1), and the reactions were performed using an iCycler iQ™ Real-Time PCR detection system (Bio-Rad Laboratories, Hercules, United States). The expression levels of the target genes were quantified relative to that of β-actin. The statistical analysis of the RT-qPCR results was performed using CFX Maestro software (Bio-Rad).

For adipocytes, proteins were extracted using RIPA buffer (Biosesang Inc.) supplemented with Xpert phosphatase inhibitor cocktail and Xpert protease inhibitor cocktail (Gendepot, Barker, TX, United States) solutions at 1:100 dilutions. For adipose tissue, protein extraction was performed using PRO-PREP™ (iNtRON Biotechnology, Seoul, Korea). Protein concentrations were adjusted to 30–60 μg using the Bradford assay (Bio-Rad), followed by the addition of 5X SDS-loading buffer (Biosesang Inc.) and boiling at 100 °C for 10 min. To separate proteins based on size, the boiled samples were loaded onto an SDS-polyacrylamide gel and subjected to electrophoresis. The separated proteins were then transferred onto a nitrocellulose membrane. To prevent the nonspecific binding of antibodies, the membrane was incubated in a blocking buffer (TBS-T with 5% skim milk) at room temperature for 1 h with shaking. Then, the membrane was incubated with the primary antibody overnight at 4 °C with shaking. The following day, the primary antibody was removed, and the membrane was washed three times in TBS-T for 5 min with shaking. The appropriate secondary antibody was then applied, and the membrane was incubated at room temperature for 1 h with shaking. The information on the antibodies used for the Western blot is provided in Supplementary Table S2. The secondary antibody was then removed, and the membrane was washed three times in TBS-T for 5 min with shaking. For band detection, the enhanced chemiluminescence (ECL) detection kit (GE Healthcare, Buckinghamshire, United Kingdom) was applied to the membrane, and images were captured using a FUSION Solo detector (Vilber Lourmat, Collégien, France). Band intensities were quantified with ImageJ software using the β-actin band’s intensity for normalization.

The amount of TNFα secreted by cells was measured in the culture medium from stimulated RAW264.7 macrophages. The assay was performed according to the manufacturer’s protocol using the Mouse TNFα ELISA Kit–Quantikine (R&D Systems, Oxon, United Kingdom). Briefly, the cell supernatant was diluted 50-fold with Calibrator Diluent. The diluted samples were then added to an antibody-coated enzyme-linked immunosorbent assay (ELISA) plate and incubated in darkness at room temperature for 2 h with shaking. After incubation, the plate was washed with washing buffer, and the conjugate was added for further reaction. After another washing step, Substrate Solution was added, and the plate was incubated in darkness for 30 min with shaking. The reaction was then terminated by adding Stop Solution, and absorbance was measured at 450 nm using an ELISA reader (Tecan).

Male C57BL/6 mice were purchased at 6 weeks of age from HyoChang Science (Daegu, Korea). Before experimentation, the mice underwent a 1-week acclimation period. For the experiment, the mice were randomly divided into four groups: (1) a normal diet group (ND group), (2) a high-fat diet group (HFD group), (3) a group administered an HFD and 10 mg/kg/day of NJFE (NJFE10 group), and (4) a group administered an HFD and 20 mg/kg/day of NJFE (NJFE20 group). To induce obesity, the HFD (a rodent diet with 60 kcal% fat; Research Diets, NJ, United States) was provided for a total of 12 weeks. The composition of the HFD by weight was as follows: 34.89% fat, 26.23% protein, 25.56% carbohydrate, 6.46% fiber, and 6.46% minerals (Licholai et al., 2018). The ND and HFD group mice received PBS as a vehicle control, while the NJFE10 and NJFE20 group mice were orally administered NJFE diluted in PBS every other day. Body weight and food intake were measured every other day. At the end of the 12-week period, tissue and blood samples were collected for analysis. All animal experiments were conducted in accordance with the ethical guidelines of Kyungpook National University (Daegu, Korea) and were approved by the Institutional Animal Care and Use Committee (Approval No. KNU-2023-0208).

Epididymal and inguinal white adipose tissues (eWAT and iWAT) were fixed in 4% paraformaldehyde overnight. The fixed tissues were embedded in paraffin and sectioned at a thickness of 5 μm. Sections were then stained with hematoxylin and eosin (H&E). Stained adipose tissues were observed and imaged using a microscope (Leica Camera AG, Wetzlar, Hesse, Germany). The area (μm²) of lipid droplets in the images was quantified using the Adiposoft plug-in in ImageJ.

Adipose tissue (eWAT) sections were deparaffinized, rehydrated through a graded ethanol series, and subjected to antigen retrieval in 10 mM citrate buffer (pH 6.0). The sections were then incubated overnight at 4 °C with anti-F4/80 antibody (macrophage marker; Cell Signaling Technology, MA, United States) diluted 1:200 in blocking solution. The following day, slides were washed with PBS and incubated with a secondary antibody for 1 h at room temperature. The staining signal was visualized using a DAB substrate, and images were captured using a light microscope (Leica Camera AG).

Mice underwent an intraperitoneal glucose tolerance test (IP-GTT) during week 11 of the in vivo experiment to measure blood glucose levels. The day before the experiment, the mice fasted for at least 12 h. On the following day, all mice received an intraperitoneal injection of D-glucose at a dose of 1 g/kg at the same time, and blood glucose levels were measured 15, 30, 60, 90, 120 min post-injection using an Accu-Check EZ glucose monitor (Roche Molecular Biochemicals, IN, United States) via the tail vein.

Blood was collected from mice via retro-orbital bleeding, and the samples were centrifuged at 3,500 rpm for 20 min at 4 °C to separate the plasma. The levels of glutamate oxaloacetate transaminase (GOT), glutamate pyruvate transaminase (GPT), blood urea nitrogen (BUN), creatinine (CREA), total cholesterol (T-CHO), high-density lipoprotein cholesterol (HDL-C), and low-density lipoprotein cholesterol (LDL-C) in the plasma were measured using an Olympus AU400 analyzer (Olympus Optical, Tokyo, Japan).

To stain mitochondria, 3T3-L1 adipocytes were incubated with MitoTracker^® Orange (Thermo Fisher Scientific, Waltham, MA, United States). First, the dye was dissolved in DMSO to prepare a 1 mM stock solution. For staining, the stock solution was diluted in pre-warmed serum-free DMEM to a final working concentration of 25 nM. Cells in a six-well plate were washed twice with PBS, and 1 mL of the MitoTracker^® working solution was added to each well. The plates were then incubated for 30 min at 37 °C in a humidified CO₂ incubator. After incubation, the cells were washed once with PBS and fixed with 4% paraformaldehyde (PFA) for 10 min at 37 °C. Following an additional PBS wash, stained cells were visualized using a fluorescence microscope (Leica Camera AG).

NJFE detection was performed using an Alliance e2695 Separation Module (Waters Corporation, Milford, MA, United States) equipped with a Waters 2489 UV/Vis detector and a QDa mass detector (Waters Corporation, Milford, MA, United States). Chromatographic separation was carried out using an Acquity UPLC C18 column (250 mm × 3.0 mm, 5 μm particle size). Ammonium formate water was used as solvent A, and 0.1% formic acid in methanol was used as solvent B, with a flow rate of 0.6 mL/min. The gradient program consisted of an initial equilibration at 95% solvent A for 10 min, followed by a linear decrease of solvent A from 95% to 0% over 5–25 min, maintained at 0% solvent A from 25 to 30 min, then increased back to 95% solvent A between 30 and 30.01 min, and finally re-equilibrated at 95% solvent A from 30.01 to 35 min. The NJFE sample was injected at a concentration of 10 ppm. Mass spectrometry was performed in negative electrospray ionization mode, scanning from 50 to 800 m/z. The probe temperature was maintained at 400 °C, and the source temperature was set to 120 °C. Collision energy was gradually increased from 10 to 40 V, and the capillary voltage in positive mode was set to 0.8 kV. The LC-MS analysis results and detailed conditions are shown in Figure 7.

All experiments were performed with three technical and three biological replicates. Data are presented as the mean ± the standard deviation for in vitro experiments and as the mean ± the standard error of the mean for in vivo experiments. Statistical significance was determined using one-way ANOVAs in IBM SPSS Statistics 27 and GraphPad Prism 9 (GraphPad Software, San Diego, CA, United States). A p-value of less than 0.05 was considered statistically significant.

We used a mouse model of obesity induced by a 12-week HFD to evaluate the anti-obesity effects of NJFE. Mice were orally administered by NJFE along with HFD. Mice were randomly assigned to groups administered an ND (Normal diet), an HFD (High-fat diet), an HFD plus 10 mg/kg/day of NJFE or an HFD plus 20 mg/kg/day of NJFE (Figure 1A). To confirm that the doses used in this study did not induce in vivo toxicity, we evaluated changes in the weights of the liver, spleen, and kidneys, as well as systemic toxicity markers. NJFE administration at all tested doses did not cause significant alterations in organ weights. In addition, serum biochemical analyses showed no significant changes in the hepatic toxicity markers GOT (aspartate aminotransferase) and GPT (alanine aminotransferase) or the renal toxicity markers BUN (blood urea nitrogen) and CREA (creatinine) compared with the control group. These results indicate that the doses of NJFE used in this study did not cause systemic toxicity in mice (Supplementary Figures S1A,B). However, the NJFE treatments significantly inhibited the accumulation of both subcutaneous and visceral fat induced by the HFD, as observed visually. Furthermore, the HFD-induced expansion of adipose tissue, including iWAT and eWAT, was also suppressed, and it was confirmed that hepatic fat accumulation induced by the HFD was significantly reduced by the NJFE treatments (Figures 1B,C). Compared to the HFD group, the NJFE treatments produced a significant weight reduction effect starting in the fourth week, and the final body weight at 12 weeks was significantly lower (Figures 1D,E). To accurately assess obesity in rodents, we calculated Lee’s Index, which is based on body weight and length, and Lee’s index was significantly lower in the NJFE treatment groups than in the HFD group (Figure 1F). In addition to weight reduction, NJFE treatments significantly decreased the weights of iWAT, eWAT, and rWAT (retroperitoneal WAT) (Figure 1G). This led to a reduction in adiposity, as reflected by decreases in the adipose tissue weights of mice relative to their body weights (Figure 1H). The weight loss effect of NJFE in obese mice was not due to changes in food intake (Supplementary Figure S1C). Therefore, the results of this study suggest that NJFE inhibits weight gain and adipose tissue expansion induced by HFD.

We investigated whether NJFE has effects beyond weight loss and reductions in adipose tissue mass. Specifically, whether it has the potential to improve obesity-induced adipocyte hypertrophy, dyslipidemia, and insulin resistance. Compared to the HFD group, the NJFE treatment groups showed significant reductions in lipid droplet area in both iWAT and eWAT (Figures 2A,B). Additionally, we observed that the serum levels of T-CHO, LDL-C, and HDL-C were lower in the NJFE treatment groups than in the HFD group (Figure 2C). To determine whether the decrease in HDL-C is due to the reduction in T-CHO, we analyzed the LDL/HDL ratio. The LDL/HDL ratio is commonly used as an indicator of cardiovascular disease risk and effectively quantifies the balance between good cholesterol and bad cholesterol in the blood. We observed a significant decrease in the LDL/HDL ratio in the NJFE-treated groups compared to the HFD group (Figure 2D). Similarly, the NJFE-treated groups also showed a significant decrease in serum triglyceride levels (Figure 2E). Regarding insulin resistance and glucose intolerance, when glucose was administered, mice in the NJFE-treated groups began to recover from the glucose-induced blood sugar increase starting 30–60 min after administration, whereas HFD-group mice did not start to recover for up to 60 min after administration. The difference in blood sugar reduction between the NJFE 20 0 group and the HFD group became significant starting at 30 min. A significant difference in blood glucose levels between the NJFE 10 group and the HFD group began at 60 min (Figures 2F,G). We also observed that the basal blood glucose levels after fasting were significantly lower in the NJFE-treated groups than in the HFD group (Figure 2H). In summary, these results suggest that NJFE significantly improves the adipocyte hypertrophy, dyslipidemia, insulin resistance, and glucose intolerance caused by diet-induced obesity.

Building on our previous finding that NJFE inhibits HFD-induced obesity and mitigates adipocyte hypertrophy, dyslipidemia, and glucose intolerance, we examined whether NJFE also alleviates obesity-induced adipose tissue inflammation. The protein F4/80 is specifically expressed in tissue-resident macrophages and is used as a marker to assess the presence and activation status of macrophages in adipose tissue. Obesity is characterized by a chronic inflammatory state in adipose tissue in which recruited adipose tissue macrophages induce inflammation and act as mediators of insulin resistance (Lumeng et al., 2007). Therefore, we first investigated whether NJFE inhibits macrophage recruitment in the inflammatory environment induced by obesity by examining F4/80 expression in eWAT. As confirmed by immunohistochemistry, the HFD induced elevated F4/80 protein expression, which was reduced by the NJFE treatments, and this reduction was consistent with decreases in F4/80 gene expression (Figures 3A,B). This led to a decrease in Tnfα expression in the NJFE treatment groups compared to the HFD group (Figure 3C). Additionally, we analyzed the NJFE treatments’ effects on adipogenesis markers and the phosphorylation levels of NF-κB in eWAT. The transcription factors PPARγ and C/EBPα are well-known master regulators involved in adipocyte differentiation and lipid synthesis, and decreases in their expression suggest a reduction in adipogenesis (Rosen, 2005). The NF-κB pathway is a prominent mechanism that induces inflammation and is a strong mediator of insulin resistance (Shoelson et al., 2003). We observed increased mRNA and protein expression levels of PPARγ and C/EBPα in HFD-fed obese mice, and this increase was significantly reduced in NJFE treatment-group mice (Figures 3D–G). At the same time, HFD-induced phosphorylation of NF-κB was also significantly decreased by the NJFE treatments (Figures 3E,H). Based on these findings, we propose that NJFE not only alleviates obesity through the inhibition of adipogenesis-related gene expression but also suppresses obesity-induced macrophage recruitment and inflammation in adipose tissue.

To assess the reproducibility of NJFE’s in vivo anti-obesity and obesity complication-ameliorating effects, we conducted in vitro experiments using 3T3-L1 adipocytes. Adipocyte differentiation was conducted over a total of 8 days, following the induction protocol shown in Figure 4A. NJFE showed no cytotoxicity up to 32 μg/mL. Therefore, we used the lowest effective concentrations, 4 and 8 μg/mL, in adipocyte experiments (Figure 4B). Treating cells with NJFE throughout the entire differentiation period led to a dose-dependent decrease in adipocyte differentiation and lipid accumulation (Figures 4C,D). To investigate whether this reduction was due to decreases in the expression of adipogenesis-related genes, we examined the protein levels of two key transcription factors regulating adipogenesis, PPARγ and C/EBPα, and the lipid accumulation marker adiponectin. The expression of these three proteins decreased in a dose-dependent manner with NJFE treatment. The IC₅₀ values for PPARγ and C/EBPα were 5.496 and 4.768 μg/mL, respectively (Figures 4E,F). Additionally, we analyzed the gene expression levels of four transcription factors, two that promote adipocyte differentiation, Pparγ and C/ebpα, and two that inhibit it, Chop10 and Gata2. Chop10 suppresses adipocyte differentiation by inactivating C/EBPβ (Huang et al., 2008), while Gata2 acts as a negative regulator of adipogenesis by interacting with and repressing PPARγ (Tong et al., 2000). The NJFE treatments reduced Pparγ and C/ebpα expression in a dose-dependent manner while increasing Chop10 and Gata2 expression. Beyond its regulation of transcription factors related to adipocyte differentiation, NJFE also significantly reduced the expression of the adipocyte maturation and lipid accumulation marker genes Ap2, Adipoq, Hsl, and Lpl (Figures 4G,H). In addition, we examined IRβ and IRS1 as upstream targets of NJFE. These molecules are key metabolites of the insulin signaling pathway and play a crucial role in adipocyte differentiation by mediating the insulin-induced expression of major adipogenic transcription factors such as PPARγ and C/EBPα (Miki et al., 2001). 8 μg/mL of NJFE treatment significantly reduced the expression of both IRβ and IRS1 (Supplementary Figure S4A). Therefore, the downregulation of IR and IRS1 may serve as a molecular mechanism by which NJFE inhibits adipocyte differentiation. Collectively, these findings suggest that NJFE inhibits adipocyte differentiation and lipid accumulation by modulating key transcription factors involved in adipogenesis.

After confirming that NJFE inhibits adipocyte differentiation, we investigated at which stage of adipocyte differentiation NJFE exerts its strongest inhibitory effect. Adipocyte differentiation was divided into three stages: early (0–2 days), middle (2–4 days), and late (4–6 days). Accordingly, NJFE was administered at various points corresponding to all combinations of these stages (Figure 4A; Supplementary Figure S2A). Lipid accumulation was significantly reduced when the NJFE was applied during the early differentiation stages. However, administration in the middle or late stages resulted in lower reductions in lipid accumulation if NJFE was not applied in the early stage, regardless of the treatment duration (Supplementary Figures S2B,C). Furthermore, under the same experimental design, we investigated the mRNA expression of Pparγ. The expression of Pparγ was significantly more reduced in the groups treated with NJFE during the early stage (B,C,D,H) than in those not treated during the early stage (E,F,G). Therefore, this result supports the ORO staining data (Supplementary Figure S2D).

The early stage of adipocyte differentiation is the most critical, as it involves the regulation of the cell cycle through the mitotic clonal expansion of preadipocytes, initiating irreversible differentiation (Audano et al., 2022). Before differentiation induction, most preadipocytes that reach confluence remain in the G1 phase and cease growth. However, when treated with the differentiation inducer MDI, the cells re-enter the cell cycle and undergo one or two rapid divisions, a process known as mitotic clonal expansion (MCE). Previous studies have shown that MCE begins approximately 16 h after MDI treatment (Lee et al., 2023). In the early stage of adipocyte differentiation, C/EBPβ and C/EBPδ function as key transcription factors (Clarkson et al., 1995). To evaluate the effect of NJFE at this stage, 3T3-L1 preadipocytes were induced to differentiate for 16 h while simultaneously treated with NJFE. Subsequent analysis revealed that NJFE treatment significantly reduced the expression levels of C/ebpβ and C/ebpδ compared to the WC (Supplementary Figure S2E). In terms of the cell cycle, pre-differentiated cells (NC) were largely distributed in the G1 phase, and MDI treatment decreased the G1 phase distribution and increased the S and G2 phase distributions. However, when NJFE was treated, the G1 phase distribution increased and the S and G2 phase distributions decreased compared to WC, and this was dose-dependent. Therefore, NJFE delays the onset of MCE, which occurs when adipocyte differentiation is initiated (Supplementary Figures S2F,G). In conclusion, these findings highlight the anti-obesity effect of NJFE, which stems from its robust inhibition of the initial gate-keeping steps of adipocyte differentiation, thereby preventing fat cell hypertrophy.

Adipose tissue, being a major site of inflammation in obese individuals, should be considered not only as a storage reservoir for lipids but also as an immune organ. Inflammation within adipose tissue is driven by adipose tissue-associated macrophages, and the shift towards an inflammatory environment leads to insulin resistance (O'Rourke, 2009). Therefore, we aimed to confirm that NJFE alleviates macrophage-derived inflammation. The NJFE did not exhibit cytotoxicity up to a concentration of 32 μg/mL in RAW264.7 macrophages (Figure 5A). In subsequent tests, NJFE treatments 4–16 μg/mL reduced the levels of nitric oxide and TNFα released by lipopolysaccharide (LPS)-activated macrophages (Figures 5B,C). Additionally, the expression of Tnfα and Il-6, which increased upon LPS treatment, was decreased by the NJFE treatments in a dose-dependent manner. We also evaluated the effect of NJFE on the expression of the proteins iNOS and COX2, which were upregulated in the macrophages upon LPS-induced inflammation. The NJFE treatments reduced both iNOS and COX2 expressions in a dose-dependent manner. Taken together, these results are evidence that NJFE alleviates LPS-induced macrophage inflammation and the release of inflammatory cytokines by macrophages in vitro.

Based on our findings that the NJFE treatments inhibited macrophage activation and macrophage-induced inflammatory responses (Figure 5) and alleviated lipogenesis-induced inflammation in adipocytes (Figure 6A), we examined whether NJFE could suppress inflammation induced by activated macrophages in adipocytes within an obese environment. In an obese environment, serum levels of free fatty acids, cholesterol, and LPSs increase, leading to a rise in macrophage numbers, which can comprise up to 40% of the cells in adipose tissue and promote polarization to the M1 phenotype, triggering an inflammatory response (Castoldi et al., 2016; Weisberg et al., 2003). This occurs because elevated serum LPS levels, like those seen in obesity, are associated with changes in the gut microbiota. As LPSs are metabolites of the cell walls of Gram-negative bacteria, they act as a key factor in activating macrophages and promoting inflammation (Caricilli et al., 2011).

Based on this, we induced RAW264.7 macrophage activation with LPS, isolated the resulting culture medium (RAW-CM), and then used it to treat 3T3-L1 adipocytes to mimic the obesity-induced inflammatory environment in adipose tissue (Figure 6B). The RAW-CM treatment increased NF-κB activation in adipocytes, but this activation was attenuated when treated with NJFE. Additionally, NJFE treatment mitigated RAW-CM-induced reductions in adiponectin expression in adipocytes (Figures 6C–E). Adiponectin, a hormone derived from adipose tissue, plays a protective role by suppressing the development of obesity-related diseases, and its expression is reduced in obesity (Ouchi and Walsh, 2007). Moreover, NJFE alleviated RAW-CM-induced increases in the expression of genes encoding inflammatory cytokines, such as Il-6, Tnfα, and Mcp1, in adipocytes. Notably, Mcp1, primarily produced by macrophages, is closely associated with macrophage infiltration into adipose tissue in the obese environment (Kanda et al., 2006). In summary, NJFE not only alleviates lipogenesis-induced inflammation in adipocytes but also effectively suppresses the inflammatory response induced by obesity-activated macrophages in adipocytes.

Obesity is characterized by increased oxidative stress and exacerbated inflammation, accompanied by immune cell infiltration into adipocytes. These changes lead to the impairment of the mitochondrial electron transport chain and a reduction in mitochondrial biogenesis. This obesity-induced mitochondrial dysfunction, in turn, results in metabolic alterations in fatty acid oxidation and lipolysis as well as insulin resistance (de Mello et al., 2018). Therefore, if NJFE can improve mitochondrial function, it may provide additional benefits in alleviating obesity and its complications. Compared to preadipocytes (NC), adipocytes subjected to lipogenesis for a total of 8 days (WC) exhibited reduced mitochondrial biogenesis due to the inflammation intrinsically caused by lipid synthesis and accumulation. However, when NJFE was administered during differentiation, mitochondrial biogenesis was improved (Supplementary Figures S3A,B). Additionally, given that CPT1 (Carnitine Palmitoyltransferase 1) is a key regulator of fatty acid oxidation in mitochondria, we examined its expression. As with mitochondrial biogenesis, CPT1 expression was reduced by lipogenesis, but NJFE treatment during differentiation restored it (Supplementary Figure S3C). Collectively, our findings suggest that NJFE not only alleviates obesity and obesity-induced inflammation in adipose tissue but also enhances mitochondrial function in adipocytes, potentially offering additional therapeutic benefits for obesity and obesity-related complications.

The presence of kaempferol derivatives in Neoshirakia japonica (Siebold & Zucc.) Esser [Euphorbiaceae] has been previously reported in the literature (Zhao et al., 2021). Kaempferol was tentatively identified in NJFE based on mass spectral data obtained in this study by comparison with an LC-MS spectral library and previously reported metabolites isolated from Neoshirakia japonica (Siebold & Zucc.) Esser [Euphorbiaceae] (Jiang and Gates, 2024; March and Miao, 2004; Zhao et al., 2021) (Figures 7A,B). Parent ion analysis using LC-MS revealed a prominent signal corresponding to the free form of kaempferol ([M-H]^-, m/z 285.06), which differs from the kaempferol derivatives commonly reported in the literature. It has been documented that kaempferol glycosides can be hydrolyzed into their aglycone forms under certain extraction conditions or due to specific sample characteristics, though it is also possible that the free form originally existed in the extract (Biesaga, 2011; Nuutila et al., 2002). The concentration of kaempferol in NJFE (10 ppm) was determined to be 2.316 μg/mL. Kaempferol has been reported to exhibit anti-obesity and anti-inflammatory activities, supporting the relevance of our findings (Alam et al., 2020; Bian et al., 2022). These findings suggest that kaempferol may be one of the bioactive metabolites contributing to the observed anti-obesity and anti-inflammatory effects of NJFE. The chemical fingerprint established here provides a foundational understanding of the bioactive metabolites contributing to the anti-obesity and anti-inflammatory effects demonstrated by NJFE.