Fibroblast Growth Factor 6 Promotes Adipocyte Progenitor Cell Proliferation for Adipose Tissue Homeostasis

The persistent excess intake of calories leads to the expansion of white adipose tissue (WAT), resulting in obesity (1,2). In individuals with obesity, WAT expands via two processes: enlargement of existing adipocytes (hypertrophy) and recruitment of new adipocytes (hyperplasia) (3,4). Adipocyte hypertrophy can trigger cell hypoxia and dysfunction, which are accompanied by chronic inflammation and fibrosis (5). The de novo differentiation of hyperplastic adipocytes from adipocyte progenitor cells (APCs) is often accompanied by the reduction of fibrosis and inflammation and improvement in insulin sensitivity (1,2,6). Emerging evidence suggests that the elevation of the adipogenic potential of APCs helps preserve metabolic health and improves glucose metabolism in individuals with obesity and/or during the aging process (7–11). Continuous adipogenesis and its maintenance depend on the presence of sufficient APCs, whereas obesity and aging are associated with a decline in the hyperplastic potential of APCs to generate new adipocytes and insufficiency of the APC pool (12,13).

Previous studies using fluorescence-activated cell sorting and genetic lineage–tracing strategies have established that platelet-derived growth factor receptor α (PDGFRα) and PDGFRβ play a key role in APC pool maintenance and adipose tissue homeostasis (14). It has been reported that the major adipocytes of inguinal WAT (iWAT) are derived from PDGFRα⁺ adipocyte precursors (15). The PDGFRβ⁺ adipocyte precursors contribute to adipocyte hyperplasia upon high-fat diet (HFD) consumption that protects against pathological gonadal WAT (gWAT) expansion (16,17). Single-cell RNA sequencing (scRNA-seq) has resolved mouse stromal vascular fraction (SVF) heterogeneity, and the APCs have been further subdivided into distinct subpopulations according to their specific functions. Using this approach, Merrick et al. (18) identified the DPP4⁺ interstitial progenitors that produce ICAM1⁺ and CD142⁺ preadipocytes in iWAT. Importantly, Schwalie et al. (19) revealed that LIN⁻SCA1⁺CD142⁺ adipocyte precursor populations in iWAT can suppress adipocyte formation in vitro and in vivo. scRNA-seq has also been used to specifically characterize the PDGFRβ⁺ APC pool in gWAT. The LY6C⁻CD9⁻PDGFRβ⁺ subpopulation represents highly adipogenic APCs, whereas the LY6C⁺PDGFRβ⁺ subpopulation shows profibrogenic and proinflammatory phenotypes, and these APCs are termed fibroinflammatory progenitors (20). Interestingly, Yasuo Oguri et al. (21) performed scRNA-seq analysis of LIN⁻ stromal cells and demonstrated that CD81 is a surface marker of beige APCs and controls beige APC proliferation and whole-body energy homeostasis.

Meanwhile, the orphan nuclear receptor NR4A1 reportedly regulates APC quiescence. Nr4a1 expression deficiency in APCs is associated with higher proliferative and adipogenic capacities, and Nr4a1^−/− APC transplantation was shown to improve insulin sensitivity in obese mice (11). Therefore, elucidating the regulatory mechanism underlying APC proliferation at the molecular level is vital to understanding fat biology and developing new approaches for treating metabolic diseases. However, the key regulators of APC proliferation are still poorly understood.

The fibroblast growth factor (FGF) family is composed of 15 paracrine, three endocrine, and four intracellular proteins. Paracrine and endocrine FGFs interact with four signaling tyrosine kinase FGF receptors (FGFR1–4) that have been implicated in a broad range of physiological processes, including development, wound healing, and cell proliferation and differentiation (22). Several FGFs, including FGF21, have recently been shown to play essential roles in the thermogenic functions of mature adipocytes and in the remodeling and browning of adipose tissues (23–26). However, the roles of specific FGFs in the regulation of APC proliferation and the corresponding consequences for metabolic performance in vivo remain unclear.

In the current study, via screening and molecular analysis, we confirmed that FGF6, a paracrine FGF, is the optimal candidate for regulating the proliferation of PDGFRα⁺ APCs via extracellular signal–regulated kinase (ERK) signaling. Additionally, even though iWAT-specific FGF6 overexpression in obese and aging mice led to several phenotypic differences among different mouse models depending on the different physiological and pathological states of mice, these mice consistently exhibited attenuation of adipose fibrosis, apoptosis, and hypoxia, along with improvement in glucose tolerance and insulin sensitivity. These findings highlight the vital and multifaceted functions of FGF6 in adipose metabolism. Conversely, the iWAT-specific blockade of FGF6 with neutralizing antibodies or deficiency of Fgf6 expression in mice administered an HFD impaired adipose tissue expansion and insulin resistance. These results implicate FGF6 as a specific proliferative factor that controls APC hyperplasia and may serve as a promising therapeutic target for obesity- and aging-induced metabolic dysfunction.

Eight-week-old male C57BL/6J mice were purchased from GemPharmatech Co., Ltd (Nanjing, China). Diet-induced obesity (DIO) was established by feeding male C57BL/6J mice an HFD (cat. no. D12492; Research Diets, New Brunswick, NJ) for 12 weeks. Naturally aging male C57BL/6 mice (16–18 months old) were raised in our laboratory. Eight- to 10-week-old wild-type (WT) and Fgf6 knockout (KO) male mice were purchased from GemPharmatech Co., Ltd (Nanjing, China). The mice were maintained in a specific pathogen-free animal facility in a standard humidity- and temperature-controlled environment under a 12-h light/dark cycle, with free access to food and water. The animal experimental protocol was reviewed and approved by the Institutional Animal Use and Care Committee of East China Normal University.

For recombination mouse FGF6 (rFGF6) administration, C57BL/6J mice were locally injected with rFGF6 (left) or PBS (right) into the iWAT once per day for the indicated number of days. BrdU was administered in the drinking water at 0.8 mg/mL for the indicated number of days. We used the Adipoq promoter to induce adipocyte-specific adeno-associated virus (AAV) 2/9–mediated gene transfer for FGF6 or GFP overexpression (Hanbio Biotechnology Co., Ltd, Shanghai, China). Mice were anesthetized by isoflurane anesthesia (5% induction and 2% maintenance). AAVs were directly injected into the iWAT on both sides (2 × 10¹⁰ plaque-forming units for the prevention model and 4 × 10¹⁰ plaque-forming units for DIO and aging models). For the prevention model, mice were fed a chow diet (CD) for 2 weeks after AAV injection and then fed an HFD for 10 weeks. For neutralizing antibody delivery, the mice were anesthetized using isoflurane anesthesia (5% induction and 2% maintenance) before the FGF6-neutralizing antibodies (cat. no. MAB238; R&D Systems) or immunoglobulin G (IgG) (cat. no. MAB002; R&D Systems) was directly injected into the iWAT on both sides of the mice (1 μg per side) once every 2 days, after which the mice were fed an HFD for 12 weeks.

Eighteen FGFs were amplified from human cDNA, fused with an HA tag at the C terminal, and directly inserted into the pCDH-CMV-MCS-EF1-Puro vector (Public Protein/Plasmid Library, Nanjing, China). The primers used are listed in Supplementary Table 1. For transfection, HEK293T cells were cultured in a six-well plate in high-glucose DMEM supplemented with 10% FBS and 1% penicillin/streptomycin, and plasmids (2 μg per well) were transfected using EZ Trans (Life-iLab, Shanghai, China). After 12 h, the medium was replaced with fresh 10% FBS–supplemented high-glucose DMEM complete medium. The supernatants were collected 36 h later, filtered using a PVDF filter (Merck Millipore, Burlington, MA), and applied to the C3H10T1/2 cell cultures.

Mouse fat mass and lean mass were measured once per week via a body composition analyzer using time domain nuclear magnetic resonance (AccuFat-1050; MAG-MED, Jiangsu, China). Briefly, mice were placed in a plastic tube and introduced into the AccuFat-1050 instrument. For each mouse, the process of body composition determination required ∼90 s, and the fat mass and lean mass were expressed as functions of body weight.

C3H10T1/2 cells (American Type Culture Collection, Manassas, VA) were cultured in high-glucose DMEM supplemented with 10% FBS and 1% penicillin/streptomycin at 37°C in the presence of 5% CO₂. For differentiation, C3H10T1/2 cells were incubated in the culture medium for 2 days (day 0); transferred to and cultured in medium supplemented with 5 μg/mL insulin, 0.5 mmol/L 3-isobutyl-1-methylxanthine, 1 μmol/L dexamethasone, and 10 μmol/L troglitazone for 2 days (day 2); and maintained in the culture medium supplemented with 5 μg/mL insulin for 6 days (the medium was changed every 2 days). rFGF6 treatment was performed on day 0, 4, or 8. For FGF screening, the medium of C3H10T1/2 cells was replaced with one of the 19 supernatants (FGF1–10 and 16–23 and negative control) plus maintenance medium (2:1) and cultured for 24 h. RNA extraction and Oil Red O staining were performed in 24-well plates, and cell viability determination was performed in 48-well plates. The treatment for each group was repeated in triplicate.

Murine SVF cells and adipocytes were isolated from iWAT and subjected to primary culture as described previously (27), with some modifications to the process. The iWAT was carefully excised and rinsed twice with cold PBS and minced and digested in a buffer supplemented with 1 mg/mL collagenase I, 0.123 mol/L NaCl, 5 mmol/L KCl, 1.3 mmol/L CaCl₂, 5 mmol/L glucose, 100 mmol/L HEPES, and 4% BSA at 37°C for 30 min. The digested iWAT pieces were filtered through a 100-μm nylon mesh and centrifuged (150g for 5 min) at room temperature. Floating mature adipocytes were collected and washed in isolation medium without collagenase for RNA and protein isolation. The cell pellets were resuspended in red blood cell lysis buffer for 5 min, centrifuged, and washed twice with the culture medium (DMEM supplemented with 20% FBS and 1% penicillin/streptomycin). The harvested cell pellets were resuspended in the culture medium, which was refreshed daily. For differentiation, confluent primary SVF cells were stimulated with 6 μg/mL insulin, 0.5 mmol/L 3-isobutyl-1-methylxanthine, 10 μmol/L dexamethasone, and 10 μmol/L troglitazone for 2 days and maintained in 6 μg/mL insulin for another 6 days. For palmitic acid (PA) treatment, differentiated primary SVF cells from the iWAT of WT and Fgf6 KO mice were treated with PA (0.5 mmol/L) for 24 h, and the cells were collected for RNA and protein isolation.

To test tolerance to cold exposure, mice were individually housed at 4°C without bedding and with free access to food and water. The core body temperature of the mice was measured using a rectal thermometer (TH-5; Braintree Scientific) at the indicated time points.

The metabolic rate was assessed by indirect calorimetry in an open-circuit Oxymax chamber with a comprehensive laboratory animal monitoring system (Columbus Instruments). After 10 weeks of HFD administration, mice were housed individually and maintained at 23°C under a 12-h light/dark cycle with free access to food and water.

For the glucose tolerance test (GTT), mice were fasted for 16 h and injected intraperitoneally with a glucose solution in saline (1.5 g/kg body weight). For the insulin tolerance test (ITT), the mice received an intraperitoneal injection of insulin (1.25 units/kg body weight). The blood glucose levels were measured from the tail blood before and 15, 30, 60, 90, and 120 min after glucose and insulin injections.

For Oil Red O staining, cells were washed twice with PBS and fixed with 10% formalin for 10 min at room temperature. Next, the cells were washed twice with distilled water and incubated in 60% isopropanol for 5 min. After aspirating the isopropanol, the cells were stained with freshly diluted and filtered Oil Red O solution (0.5% Oil Red O dissolved in a 3:2 isopropanol to distilled water solution) for 20 min in the dark at room temperature, rinsed three times in distilled water, and imaged. For hematoxylin-eosin, Masson trichrome, and TUNEL staining, the brown adipose tissue (BAT), iWAT, and gWAT were fixed overnight in 4% formalin, embedded in paraffin, and sectioned into sections 5 μm in thickness. The sections were stained using the corresponding kits (hematoxylin-eosin and Masson trichome staining kits, Servicebio Technology; TUNEL staining kit, Roche) according to the manufacturers’ instructions. For immunohistochemistry (IHC), sections of iWAT were pretreated with 3% H₂O₂ for 30 min to quench endogenous peroxidase activity, blocked in PBS containing 2.5% horse serum for 1 h, and treated overnight with the corresponding primary antibodies against PDGFRα (cat. no. 3174S; Cell Signaling), UCP1 (cat. no. ab209483; Abcam), COX IV (cat. no. GB11250; Servicebio Technology), F4/80 (cat. no. sc-52664; Santa Cruz Biotechnology), hypoxia-inducible factor 1α (cat. nos. NB100–105; Novus Biologicals), and α-SMA (cat. no. GB111364; Servicebio Technology) in a humidified chamber at 4°C. Sections were washed with PBS, incubated with an IgG–horseradish peroxidase (Servicebio Technology) secondary antibody for 1 h at room temperature, and visualized using a DAB kit (Servicebio Technology). Images were acquired using an Olympus microscope.

Cells or iWAT tissues were homogenized in RIPA buffer (50 mmol/L Tris-HCl, 150 mmol/L NaCl, 5 mmol/L MgCl₂, 2 mmol/L EDTA, 1 mmol/L NaF, 1% NP-40, and 0.1% SDS) containing a protease inhibitor cocktail (Beyotime). The protein samples were then quantified with a BCA kit (Beyotime), fractionated by SDS-PAGE, and transferred to nitrocellulose membranes (Merck Millipore). Membranes were blocked in 5% milk and treated overnight at 4°C with the following primary antibodies: rabbit anti-PDGFRα (cat. no. 3174S; Cell Signaling), rabbit anti–phosphorylated Akt (p-Akt) (Ser473) (cat. no. 4060S; Cell Signaling), rabbit anti-Akt (cat. no. 4685S; Cell Signaling), rabbit anti–α-SMA (cat. no. GB111364; Servicebio Technology), mouse anti–p-ERK1/2 (Thr202/Tyr204) (cat. no. 4370T; Cell Signaling), mouse anti-ERK1/2 (cat. no. sc-514302; Santa Cruz Biotechnology), mouse anti-FGF6 (cat. no. sc-374518; Santa Cruz Biotechnology), rabbit anti–p-JNK (Thr183/Tyr185), rabbit anti–p-p38 (Thr180/Tyr182) (cat. no. 9910T; Cell Signaling), rabbit anti–p-STAT3 (Tyr705) (cat. no. 9145S; Cell Signaling), rabbit anti–α-tubulin (cat. no. AF0001; Beyotime), mouse anti–β-actin (cat. no. sc-8432; Santa Cruz Biotechnology), and mouse anti-GAPDH (cat. no. ET1601; Huabio). Signals were detected using the Odyssey imaging system (LI-COR) after treatment with secondary antibodies for 1 h at room temperature.

Total RNA was extracted from tissues or cells using the RNAiso plus reagent (Takara Bio, Kusatsu, Japan) and reverse transcribed to cDNA using the PrimeScript RT Reagent Kit with gDNA Eraser (Takara Bio) according to the manufacturer’s instructions. Data analysis was performed using an SYBR green PCR mix (Vazyme, Nanjing, China) on a Light Cycler 480 (Roche). Relative quantification analysis of the gene expression data was performed using the 2^−ΔΔCt method. The relative expression results were normalized to 36B4 mRNA levels. The primers used are listed in Supplementary Table 2.

Statistical analyses were performed using Prism 7.0 software (GraphPad). Significance in two-group comparisons was determined using a two-tailed Student t test. One-way ANOVA followed by the Tukey post hoc test was used for multiple group comparisons. Correlations were examined with a nonparametric Spearman correlation test. In vitro experiments were repeated at least in triplicate; the number of mice used per experiment is shown in the figure legends. Differences or values with P < 0.05 were considered statistically significant in all experiments. Data are displayed as mean ± SEM. Significant differences among groups are indicated as *P < 0.05 and **P < 0.01.

All data generated or analyzed during this study are included in the published article (and Supplementary Material).

To identify regulators of APC proliferation among the paracrine and endocrine FGFs, we screened 18 secretory FGFs (FGF1–10 and 16–23) to investigate their role in APC proliferation by transfecting individual plasmids into 293T cells. The culture supernatant was collected, filtered, and used to treat C3H10T1/2 cells (a mesenchymal stem cell line), which exhibit the potential to differentiate into adipocytes (Supplementary Fig. 1A). The concentration of each FGF in the supernatant was confirmed by Western blotting (Supplementary Fig. 1B). The population of C3H10T1/2 cells (MD-C3) at the midstage of differentiation, which included fibroblasts, preadipocytes, and mature adipocytes, was used for the screening assay. At 24 h posttreatment with the supernatant containing the FGFs, cell viability was determined using the CCK-8 assay, and the expression of Pdgfrα, Ly6a, Pdgfrβ, Acta2, and Cd34, which are common surface markers of APCs, was measured using real-time PCR. Of note, among the 18 tested FGFs, FGF6 exerted the strongest effect on cell proliferation and specifically induced Pdgfrα and Ly6a expression (Fig. 1A and Supplementary Fig. 1D and E). In addition, the proliferative effect of FGF6 was further confirmed in undifferentiated cells (Supplementary Fig. 1C). To confirm this unique role of FGF6, we treated C3H10T1/2 cells with different doses of rFGF6 at different stages of differentiation. Consistently, rFGF6 treatment induced significant PDGFRα expression and cell viability in a dose-dependent manner, even in fully differentiated adipocytes at day 8 (Fig. 1B–F and Supplementary Fig. 2A–C). To clarify the roles of FGF6 in iWAT, we performed BrdU tracing experiments (Fig. 1G and Supplementary Figs. 3A and 4A). We found that the BrdU⁺PDGFRα⁺ APCs were significantly increased in rFGF6-treated iWAT compared with in the PBS-treated iWAT (Fig. 1H–J and Supplementary Fig. 4B), even under cold or HFD stress (Supplementary Figs. 3B and 4C). Of note, compared with CD, 1 week of HFD consumption decreased BrdU⁺PDGFRα⁺ APCs by 3.5% and 4.5% in PBS- and rFGF6-treated mice, respectively (Supplementary Fig. 4B and C). In addition, decreased expression of adipocyte markers in mature adipocytes of rFGF6-treated mice further confirmed the enhanced adipogenesis upon rFGF6 treatment (Supplementary Fig. 4D). These data indicate that FGF6 induces PDGFRα⁺ APC proliferation and enhances adipogenesis to form new adipocytes.

Given that FGF6 is a paracrine protein that exhibits regulatory roles in its local environment, we first assessed the FGF6 expression pattern and found that FGF6 was detectable in BAT and iWAT but undetectable in gWAT and was primarily expressed in fully differentiated adipocytes (Supplementary Fig. 5A–C). Because overnutrition or aging leads to APC insufficiency and adipocyte hypertrophy, we further examined FGF6 mRNA and protein levels in mature adipocytes isolated from the iWAT of obese and aging mice. Of note, compared with that in control mice, the expression of FGF6 significantly decreased, whereas adipose hypoxia and fibrosis increased (Supplementary Fig. 5D–F). These data indicate that FGF6 secreted by adipocytes may be involved in the regulation of the APC pool.

On the basis of the role of FGF6 in APC proliferation, we further investigated the intracellular signaling pathway that FGF6 induced in the process. To determine the intracellular cascade that regulates APC proliferation, we treated MD-C3 cells with rFGF6 and examined the proteins involved in FGF-related signaling pathways, including STAT3, Akt, and mitogen-activated protein kinase (22). Notably, ERK1/2 phosphorylation uniquely and significantly increased in response to rFGF6 treatment (Fig. 2A), which is consistent with findings on ERK signaling activation in cell proliferation in other systems (28,29). Consistent with this finding, PDGFRα mRNA and protein expression and ERK signaling activation were increased in mice that received a single iWAT-specific delivery of rFGF6 (Fig. 2B–D). Detailed genetic analysis showed that acute rFGF6 treatment significantly increased Ki67 expression and reduced the expression of genes associated with fibrosis and apoptosis, whereas it exerted no effect on the expression of adipocyte marker genes (Fig. 2C and Supplementary Fig. 6). Furthermore, treatment of MD-C3 cells with FR-180204, an ERK1/2 inhibitor, largely reversed the rFGF6-induced increase in Pdgfrα, Ly6a, and Ki67 mRNA levels and PDGFRα expression and ERK1/2 phosphorylation and decreased the viability of the cells (Fig. 2E–G).

A recent study demonstrated that FGF6 can regulate the transcription of Ucp1 (26). To evaluate the roles of FGF6 in beige fat formation, we first examined the expression of FGF6 upon cold stress and found that 3 days of cold exposure significantly induced the expression of FGF6 in iWAT (Supplementary Fig. 7A and B). Next, C57BL/6J mice were treated with rFGF6 or PBS and kept in normal or cold conditions, and we found that rFGF6 treatment increased UCP1 levels, which was enhanced under cold conditions (Supplementary Fig. 7C–F). However, rFGF6 treatment had no effect on Il33 expression (Supplementary Fig. 7G), which has been suggested to be a regulator of iWAT beiging (30). These results indicate that FGF6 could promote inguinal fat browning.

Of note, we used primary iWAT SVF cells isolated from Fgf6 WT and KO mice to assess the effect of FGF6 on adipocyte development. As shown in Fig. 2H–K, Fgf6 KO cells showed lower vitality than Fgf6 WT cells. Over 8 days of differentiation, we treated the differentiated cells with PA to mimic the overnutrition status. Oil Red O staining revealed that Fgf6 KO adipocytes were fewer in number, larger in volume, and contained a greater number of lipid droplets (Fig. 2I). These characteristics were accompanied by a reduction in the expression of APC markers and ERK1/2 phosphorylation and an increase in the expression of adipogenesis- and apoptosis-related genes (Fig. 2J and K). These data suggest that FGF6 induces APC proliferation via ERK signaling, whereas Fgf6 expression deficiency impairs APC proliferation and may induce adipocyte hypertrophy.

Although WAT expansion is beneficial for maintaining metabolic homeostasis in obesity, the preferential accumulation of gWAT, which is associated with central obesity, is correlated with insulin resistance and an increased risk of type 2 diabetes (31–34). In contrast, the expansion of iWAT as peripheral adipose tissue may confer protection against systemic insulin resistance (35,36). We used the Adipoq promoter to generate an adipocyte-selective AAV and injected it into iWAT to achieve local and selective FGF6 overexpression (Fig. 3A). Eight-week-old C57BL/6 mice were injected with AAV-Adipoq-FGF6 (AAV-FGF6) or AAV-Adipoq-GFP (AAV-GFP) and fed an HFD for 10 weeks. Mice with iWAT-specific FGF6 overexpression (FGF6-OE HFD mice) showed significant reduction in HFD-induced body weight and fat mass gain and improved insulin sensitivity, as indicated by better performance in GTT and ITT (Fig. 3B–E). Next, we measured energy expenditure using a comprehensive laboratory animal monitoring system and found that oxygen consumption increased in FGF6-overexpressing mice, and, more notably, the difference was enhanced under cold stress (Fig. 3F and Supplementary Fig. 8A), with no significant difference in food intake or locomotor activity (Supplementary Fig. 8B and C). Consistent with these findings, FGF6-OE HFD mice showed greater cold tolerance than control mice (Supplementary Fig. 8D), which confirmed the higher energy expenditure in FGF6-OE HFD mice. In addition, tissue weight measurements and histological examination showed that both iWAT and gWAT weights were significantly lower in FGF6-OE HFD mice, with a pronounced reduction in the volume of adipocytes and lower lipid infiltration in the liver and BAT (Fig. 3G and Supplementary Fig. 8E). Overall, these results demonstrate that HFD-fed mice with continuous iWAT-specific FGF6 overexpression showed greater energy expenditure and were protected against body weight gain and metabolic dysfunction.

Previous studies have shown that PDGFRα⁺ APCs can contribute to the formation of both beige and white adipocytes (37). In addition, FGF6 was recently shown to be a UCP1 inducer, and pretreatment with FGF6 programmed human WAT SVF cells to form beige adipocytes upon differentiation (26). We investigated whether FGF6-induced PDGFRα⁺ APC proliferation enhances the formation of beige fat. As shown in Fig. 3H and Supplementary Fig. 8F, the mRNA levels of Pdgfrα and Ucp1 were significantly higher in the iWAT of FGF6-OE HFD mice, which was further confirmed with IHC. Notably, the PGC1α and UCP1 protein levels in iWAT were significantly higher in FGF6-OE HFD mice exposed to the cold for 7 days (Fig. 3I and J). Next, we performed a comprehensive analysis of the metabolic state of iWAT using quantitative PCR, IHC, immunofluorescence analysis, and Western blotting and found that iWAT with FGF6 overexpression showed activation of ERK signaling and attenuation of HFD-induced fibrosis, apoptosis, hypoxia, and inflammation (Fig. 3K–O and Supplementary Fig. 8F–I). In addition, measurement of FGF6 protein levels in other tissues, including BAT, gWAT, and liver and muscle tissues, confirmed its localized overexpression in iWAT (Supplementary Fig. 8J). These results indicate a unique role of FGF6 in preventing HFD-induced obesity via the facilitation of PDGFRα⁺ APC proliferation and inducing of beige fat biogenesis.

To further determine the therapeutic effects of FGF6 in obesity rather than its preventative effects, 8-week-old C57BL/6 mice were fed an HFD for 12 weeks to generate the DIO mouse model. The obese mice were locally injected with AAV-Adipoq-FGF6 or AAV-Adipoq-GFP, as described in Fig. 3A, and fed an HFD for an additional 6 weeks. Although no significant difference was observed in body weight or fat mass (Fig. 4A and Supplementary Fig. 9A), 6 weeks of FGF6 treatment in iWAT significantly improved glucose tolerance and insulin sensitivity (Fig. 4B and C). Interestingly, mice with DIO with the iWAT-specific overexpression of FGF6 (FGF6-OE DIO mice) showed a significant increase in iWAT expansion along with reduction in gWAT weight, lipid accumulation in the liver and BAT, and adipocyte volume in the gWAT (Fig. 4D and E). Notably, the smaller adipocyte sizes in the iWAT of FGF6-OE DIO mice suggest the importance of FGF6 in hyperplasia, accompanied by attenuation of adipose fibrosis, apoptosis, inflammation, and hypoxia (Fig. 4F–I and Supplementary Fig. 9B–D). Of note, in addition to the increase in the levels of PDGFRα, the levels of the markers of mature adipocytes Lep, Retn, and Adipoq were significantly reduced in the iWAT of FGF6-OE DIO mice (Fig. 4J). Interestingly, we found significantly increased UCP1 protein levels, even though there was no difference at the mRNA level (Supplementary Fig. 9E). In addition, Western blot analysis of FGF6 in BAT, gWAT, and muscle and liver tissues further confirmed the specific overexpression of FGF6 in iWAT (Supplementary Fig. 9F). These results indicate that FGF6 primarily induced iWAT expansion in DIO mice by promoting adipocyte hyperplasia rather than hypertrophy.

Aging is associated with APC insufficiency and limitation of the plasticity and expandability of iWAT, which leads to visceral adiposity and ectopic lipid deposition in the liver and muscles (38–41). To examine whether FGF6 treatment improves the symptoms of metabolic disorders caused by aging, 16-month-old mice were injected with AAV-Adipoq-FGF6 or AAV-Adipoq-GFP (Fig. 3A). During the 10 weeks in which the mice were fed a CD, mice with iWAT-specific FGF6 overexpression (FGF6-OE aging mice) showed resistance to body weight and fat mass gain compared with the control mice (Fig. 5A and B). Sustained FGF6 treatment considerably improved glucose tolerance and insulin sensitivity (Fig. 5C and D). Interestingly, no difference was observed in the mass of iWAT, whereas the gWAT mass was significantly reduced in FGF6-OE aging mice (Fig. 5E). Moreover, aging mice with sustained iWAT-specific FGF6 treatment showed an amelioration of adipose hypertrophy, ectopic lipid deposition, fibrosis, apoptosis, inflammation, and hypoxia (Fig. 5F–I and Supplementary Figs. 10A and B). Given the longstanding notion that the age-dependent senescence of APCs is associated with mitochondrial dysfunction (41,42), we assessed whether FGF6-induced Pdgfrα⁺ APC proliferation improves mitochondrial function. As shown in Fig. 5J and K and Supplementary Fig. 10B–F, FGF6 overexpression significantly enhanced PDGFRα expression and induced the expression of mitochondrial genes and UCP1. Consistent with these findings, Western blot analysis showed that sustained iWAT-specific FGF6 overexpression activated ERK signaling, improved insulin-stimulated p-Akt expression, and decreased fibrosis-related α-SMA protein expression, with no leakage into other tissues (Fig. 5L and Supplementary Fig. 10G). These data suggest that the potential role of FGF6 in APC proliferation could help rescue aging-associated APC insufficiency and mitochondrial dysfunction.

Overnutrition induces APC recruitment and accelerates APC insufficiency (13). FGF6 loss-of-function models were analyzed to assess the pathological functions of FGF6 in vivo. Eight-week-old C57BL/6 mice underwent chronic treatment with an injection of an FGF6-neutralizing antibody (FGF6 Ab mice) or IgG into iWAT every 2 days and were fed an HFD for 12 weeks (Fig. 6A). After HFD feeding, FGF6 Ab–treated mice showed attenuated body weight and fat mass gain compared with control mice (Fig. 6B and C). However, the results of both the GTT and ITT were worse in FGF6 Ab–treated mice (Fig. 6D and E), indicating severe insulin resistance. Of note, FGF6 Ab–treated mice exhibited lower iWAT mass, larger adipocyte volume, and higher collagen concentration (Fig. 6F and G and Supplementary Fig. 11A). Comparison of the gene expression profiles of the iWAT in IgG- and FGF6 Ab–treated mice revealed lower Pdgfrα and Ly6a mRNA levels and greater expression of fibrosis-, apoptosis-, and inflammation-related genes, whereas no difference was observed in the expression of adipogenesis- or thermogenesis-related genes (Fig. 6H and Supplementary Fig. 11B–E). Consistent with these findings, Western blot analysis showed that the chronic iWAT-specific blockade of FGF6 expression decreased the protein levels of PDGFRα, p-Akt, and p-ERK1/2 but increased those of α-SMA (Fig. 6I).

Next, we used WT and Fgf6 KO mice, which showed no metabolic abnormalities under normal conditions (43), to investigate the role of FGF6 in mediating HFD-induced APC recruitment and proliferation. Compared with the WT mice, Fgf6 KO mice showed a substantial reduction in body weight and fat mass gain, whereas no difference was seen in lean mass in response to 18 weeks of HFD feeding (Fig. 7A–C and Supplementary Fig. 12A). However, Fgf6 KO mice exhibited significantly poorer glucose tolerance, even with a lower food intake (Fig. 7D and Supplementary Fig. 12B). Notably, the average ratio of iWAT weight (normalized to body weight) in Fgf6 KO mice was approximately 41% lower than that in WT mice; in contrast, the average ratio of gWAT increased by ∼81% (Fig. 7E and F). Histological analysis revealed adipocyte hypertrophy, with macrophage infiltration, inflammation, and fibrosis, in the iWAT and gWAT of Fgf6 KO mice (Fig. 7G and H). Consistent with this finding, detailed genetic analysis of Fgf6 KO mice revealed the downregulation of Pdgfrα and upregulation of apoptosis- and fibrosis-related genes after 18 weeks of HFD feeding (Fig. 7I). These data demonstrate that FGF6 may contribute to HFD-induced APC recruitment, whereas FGF6 expression blockade or deficiency accelerates APC insufficiency.

The ability of APCs to proliferate continuously and repopulate adipose tissues with new adipocytes is critical to maintaining metabolic homeostasis (6,12,16). Nevertheless, persistent overnutrition and aging accelerate the insufficiency of the APC pool, leading to adipocyte hypertrophy, ectopic lipid deposition, and insulin resistance (13). Therefore, understanding the mechanisms underlying APC proliferation is important for treating obesity and aging-associated metabolic dysfunction. In the current study, we demonstrated that FGF6 acts in an autocrine and/or paracrine manner to control PDGFRα⁺ APC proliferation via ERK signaling. Notably, FGF6 overexpression in iWAT ameliorated adipose fibrosis, apoptosis, and inflammation; ectopic lipid deposition; and insulin resistance in the HFD, DIO, and aging mouse models. Conversely, HFD-fed mice with chronic iWAT-specific FGF6 blockade (mediated by a neutralizing antibody) or Fgf6-null mice showed severe adipocyte hypertrophy, adipose fibrosis, inflammation, and impaired glucose tolerance.

Reportedly, several proteins of the FGF family are involved in adipose tissue expansion and affect insulin sensitivity. For instance, Fgf1-deficient mice failed to exhibit gWAT expansion, which indicated the role of the PPARγ-FGF1 axis in the regulation of adipose tissue remodeling (25). Of note, in our screening system, FGF6 exhibited stronger effects than FGF1 in inducing APC proliferation. Likewise, FGF21 has been proposed to be a regulator of subcutaneous fat expansion, and HFD-fed mice with Fgf21 KO showed reduced subcutaneous fat and greater insulin resistance (8). Although several studies have concluded that iWAT expands almost exclusively via adipocyte hypertrophy (4,44), conflicting results have been reported in other studies (45,46). These discrepant results may be attributed to the different experimental models and transgenic mice used in the different studies. Our BrdU-labeling assay results showed that 1 week of HFD consumption dramatically decreased the number of BrdU⁺PDGFRα⁺ APCs compared with the CD, indicating possible adipogenesis in iWAT upon HFD exposure. However, as a thymidine analog, BrdU is only a marker of DNA synthesis, and it can be incorporated during several non–cell-division events such as DNA repair and cell-cycle reentry (47). With the determination of a cell-cycle marker (Ki67) and APC marker (PDGFRα), we can deduce the proliferative effect of FGF6. Our study lacked genetic lineage tracing evidence to support the de novo adipogenesis in FGF6-treated iWAT; however, the hyperplastic phenotypes upon FGF6 treatment in several mouse models could help to clarify it. In addition, we also propose that the increased apoptosis may contribute to the impaired expansion of iWAT in FGF6 blockade or KO mice, in accordance with a previous study with similar phenotypes (48). Therefore, our findings demonstrate for the first time that FGF6 controls APC proliferation, induces iWAT expansion via hyperplasia, and improves insulin sensitivity in response to overnutrition or aging; these results indicate that FGF6 is a strong paracrine FGF with potential in the treatment of metabolic diseases.

Interestingly, we found several phenotypic differences, including those in body weight, fat mass, and expression of several groups of genes, among the three types of animal models used in our study. We can summarize the function of FGF6 in these models as follows: tolerance to HFD, as indicated by the prevention of body weight gain and reduction of both iWAT and gWAT mass in HFD-fed lean mice; tolerance to aging, as indicated by the prevention of body weight gain and reduction of gWAT mass in CD-fed aging mice; and adaptation to DIO, as indicated by the healthy expansion of iWAT in HFD-fed mice with DIO (Fig. 7K). According to a recent report, FGF6 can induce UCP1 expression in preadipocytes (26); our findings indicate a vital role of FGF6 in PDGFRα⁺ APC proliferation. Therefore, on the basis of the considerable differences in the physiological and pathological states of the animal models, we identified the following potential mechanisms: 1) FGF6 expression in HFD-fed lean mice helped maintain a sufficient number of APCs, induced UCP1 expression in preadipocytes, and reduced body weight and fat mass; 2) sustained FGF6 treatment ameliorated APC shortage and adipocyte hypertrophy in aging mice, because the caloric intake was normal; and 3) HFD-fed mice with DIO showed severe APC insufficiency and persistent overnutrition, which rapidly promoted the FGF6-induced differentiation of APCs into adipocytes, thus increasing the fat mass in iWAT.

With the subdivision and functional identification of the APC populations, a variety of cell subsets are also found to play a regulatory role by secreting factors. For example, a recent study revealed that DPP4⁺PDGFRβ⁺ APCs act as cold-responsive cells that regulate the beiging of iWAT through the production of interleukin-33 (30). In addition, Hua Dong et al. (49) identified RSPO2 as a functional regulator of adipogenesis, which is secreted by CD142⁺ APCs to inhibit adipogenesis and impair adipose tissue homeostasis. However, the expression of Il33 was unchanged upon rFGF6 treatment, indicating the direct regulation of Ucp1 transcription via the FGF6-PGE2-ERRA-FLII-UCP1 pathway (26).

Another interesting phenotype is the decreased gWAT mass among all three FGF6 overexpression models. As the two main sites for fat storage, iWAT and gWAT should be balanced, and the storage pressure upon overnutrition will be relieved with an improved iWAT metabolic state (hyperplasia). In addition, we found that the expression of the central obesity-associated gene, Retn (encoding Resistin) (50), was significantly decreased upon FGF6 treatment. This may be a direct mechanism for the decreased gWAT in the obesity and aging mouse models.

Even though skeletal muscle is the primary site for FGF6 expression, Fgf6 KO mice are viable and fertile and show normal skeletal muscle formation (43). In addition, evidence from FGF6 gain- or loss-of-function studies in skeletal muscles suggests that FGF6 may not be indispensable in myogenesis (51–54). Consistent with these findings, the Fgf6 KO mice did not exhibit changes in morphological characteristics under basal conditions, and no significant differences in lean mass were observed between HFD-fed WT and Fgf6 KO mice. Findings from various previous studies, along with our results, have shown that HFD feeding induces a drastic response in the adipose tissues of mice but has a limited effect on lean mass (55–58). Because FGF6 expression is low in adipose tissues and undetectable in gWAT, the primary concern is whether endogenous FGF6 expression is associated with physiological functions in adipose tissues. In addition, FGF6 is a paracrine factor that is released into the extracellular matrix and functions through paracrine signaling mechanisms (59,60). Fgf6 KO mice fed an HFD for 18 weeks showed significant impairment of BAT and iWAT expansion. In addition, Fgf6 KO mice showed a higher gWAT mass than WT mice, indicating that Fgf6 defects exert limited effects on gWAT expansion and induce excess fat storage in gWAT, while impairing iWAT expansion. These findings suggest that FGF6 is a vital regulator and plays a critical role in fat homeostasis.

Taken together, our findings indicate that FGF6 guards the APC pool and controls PDGFRα⁺ APC proliferation in adipose tissues. These data further indicate the critical role of FGF6 in preventing adipocyte hypertrophy in cases of overnutrition and aging and maintaining fat homeostasis and insulin sensitivity.

Funding. The current study was supported by grants from the National Key Research and Development Project of China (2018YFA0800402 and 2019YFA0904500), the National Natural Science Foundation of China (81900766, 81974118, 32022034, and 32071148), Shanghai Outstanding Academic Leaders (20XD1433300), the Shuguang Project (21SG11), and the Innovative Research Team of High-Level Local Universities in Shanghai (SHSMU-ZDCX20212700).

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. C.L. wrote the manuscript. C.L. and M.M. performed the experiments. C.L., X.M., and C.H. conceptualized and designed the experiments. B.X., Y.X., G.L., Y.C., D.W., and J.Q. assisted with the in vivo experiments. G.L. assisted in the genotyping of Fgf6 KO mice. J.Y. assisted in the determination of the energy expenditure of the mouse models via the comprehensive laboratory animal monitoring system. L.X., X.M., and C.H. reviewed and revised the manuscript. C.H. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.