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Acta Pharmacologica Sinica logoLink to Acta Pharmacologica Sinica
. 2024 Jan 26;45(5):988–1001. doi: 10.1038/s41401-024-01226-7

Macrophage-specific FGFR1 deletion alleviates high-fat-diet-induced liver inflammation by inhibiting the MAPKs/TNF pathways

Yan-ni Zhao 1,2,3, Zhou-di Liu 2, Tao Yan 2, Ting-xin Xu 2, Tian-yang Jin 2, Yong-sheng Jiang 1, Wei Zuo 1, Kwang Youl Lee 3,✉, Li-jiang Huang 1,✉, Yi Wang 1,4,✉
PMCID: PMC11053141  PMID: 38279043

Abstract

Non-alcoholic fatty liver disease (NAFLD) is a common metabolic disease that is substantially associated with obesity-induced chronic inflammation. Macrophage activation and macrophage-medicated inflammation play crucial roles in the development and progression of NAFLD. Furthermore, fibroblast growth factor receptor 1 (FGFR1) has been shown to be essentially involved in macrophage activation. This study investigated the role of FGFR1 in the NAFLD pathogenesis and indicated that a high-fat diet (HFD) increased p-FGFR1 levels in the mouse liver, which is associated with increased macrophage infiltration. In addition, macrophage-specific FGFR1 knockout or administration of FGFR1 inhibitor markedly protected the liver from HFD-induced lipid accumulation, fibrosis, and inflammatory responses. The mechanistic study showed that macrophage-specific FGFR1 knockout alleviated HFD-induced liver inflammation by suppressing the activation of MAPKs and TNF signaling pathways and reduced fat deposition in hepatocytes, thereby inhibiting the activation of hepatic stellate cells. In conclusion, the results of this research revealed that FGFR1 could protect the liver of HFD-fed mice by inhibiting MAPKs/TNF-mediated inflammatory responses in macrophages. Therefore, FGFR1 can be employed as a target to prevent the development and progression of NAFLD.

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Keywords: non-alcoholic fatty liver disease, fibroblast growth factor receptor 1, macrophages, MAPKs, NF-κB, hepatic stellate

Introduction

In humans, liver is the largest internal organ that plays an essential role in metabolic processes, including glucose metabolism, fatty acid metabolism, amino acid metabolism, and hormone metabolism [1]. Liver dysfunction can dysregulate these metabolisms, causing metabolic disorders and other health problems [2]. Non-alcoholic fatty liver disease (NAFLD) is a common metabolic disorder with a high global prevalence and is manifested with excessive fat accumulation in the liver, causing serious consequences such as chronic inflammation and fibrosis [3]. NAFLD induces multiple complications, such as cirrhosis and liver cancer [4]. Therefore, finding effective targets for NAFLD treatment is urgent. Recent studies have shown that NAFLD development involves macrophages [5], which are activated by fat accumulation and cell damage, resulting in the release of many inflammatory mediators such as TNF-α, IL-6, and IL-1β. These inflammatory mediators further trigger the inflammatory response of other immune cells, leading to abnormal liver lipid metabolism and promoting the occurrence and development of liver fibrosis [6, 7]. Therefore, inhibiting key inflammatory signaling proteins in macrophages may confer significant protection against NAFLD.

Fibroblast growth factor receptor 1 (FGFR1) is a membrane-bound tyrosine kinase receptor that belongs to the fibroblast growth factor receptor family and is involved in many important physiological and pathological processes [8]. Furthermore, point mutations in FGFR1 cause severe impairment in cranial, digital, and skeletal development [9], whereas its overexpression is related to breast cancer and prostate cancer [9]. Moreover, FGFR1 is associated with adipose tissue function and remodeling and has been indicated to participate in adipocyte precursor differentiation by activating PI3K/AKT and MEK/ERK signaling in obesity [10]. A study reported that high-fat diet (HFD) and ob/ob mice have elevated phosphorylation of FGFR1 in epididymal adipose tissues [11]. Disruption of FGF signaling significantly reduced the apoptosis and TNF-α induced inflammatory responses in hepatic stellate cells (HSCs) [12]. FGFR1 on the surface of macrophages can be stimulated by various growth factors, activating downstream pathways, including ERK, PI3K/AKT, and STAT3, which then stimulate the production and release of various inflammatory mediators, thereby exacerbating the inflammatory response [13–15]. However, whether FGFR1 in macrophages contributes to NAFLD and the potential underlying mechanisms are largely unknown.

This investigation evaluates the inflammation-suppression effects of FGFR1 in macrophages in a diet-induced NAFLD model, and the potential molecular mechanism underlying this phenomenon was studied. Furthermore, it provides evidence that gene knockout or pharmacological inhibition of FGFR1 can attenuate NAFLD in mice, suggesting that FGFR1 is a crucial factor in NAFLD development.

Materials and methods

Reagents

Antibody against F4/80 (cat# sc-377009) and HNF-4α (sc-374229) was purchased from Santa Cruz Biotechnology (Dallas, TX). Anti-smooth muscle actin-alpha (α-SMA, cat# ab5694) and p-FGFR1 (cat# 59194) antibodies were provided by Abcam (Cambridge, UK). FGFR1 (cat# 9740), IκBα (cat# 4812), phospho-NF-κB p65 (cat# 3033), and NF-κB p65 (cat# 8242) antibodies were purchased from Cell Signaling Technology (Danvers, MA). The Collagen Type I (Col-1, cat# 14695-1-AP), TGF-β (cat# 21898-1-AP), CoraLite488-conjugated Goat Anti-Rabbit IgG(H + L) (cat# SA00013-2) and CoraLite594-conjugated Goat Anti-Mouse IgG (H + L) (cat# SA00013-3) antibodies were acquired from Proteintech (Rosemont, IL). GAPDH antibody (cat# AB-P-R001) was from Hangzhou Goodhere Biotechnology (Hangzhou, China). DyLight 488-conjugated goat anti-mouse IgG (cat# A23210) and DyLight 549-conjugated goat anti-rabbit IgG (cat# A23420) antibodies were purchased from Abbkine (Wuhan, China).

AZD4547, a specific FGFR1 inhibitor [16], was acquired from Shanghai Kai Yu Pharmatech Technology Co., Ltd. (Shanghai, China). Pierce ECL Western Blotting substrate and diaminobenzidine (DAB) were purchased from Thermo Fisher (Waltham, MA). Assay kits for total cholesterol (TCH; cat# A111-1-1), triglyceride (TG; cat# A110-1-1), low-density lipoprotein-cholesterol (LDL-C; cat# A113-1-1), alanine aminotransferase (ALT; cat# C009-2-1), and aspartate aminotransferase (AST; cat# C010-2-1) were purchased from Nanjing Jiancheng Bioengineering Institute (Nanjing, China). ELISA kits for mouse TNF-α (cat# 88-7324-76) and IL-6 (cat# 88-7064-76) cytokine were acquired from Invitrogen (Waltham, MA), whereas that for TGF-β (cat# 70-EK981-96) was obtained from Multi Sciences (Hangzhou, China). Masson’s Trichrome Stain Kit (cat# 1340), hematoxylin and eosin (H&E) staining kit (cat# G1120), Sirius red (cat# S8060), and Oil-Red O staining solution (cat# G1261) were purchased from Solarbio Life Sciences (Beijing, China). Sodium palmitate (PA; cat# P9767-5G) and bovine serum albumin (BSA; cat# A1933) were purchased from Sigma (Louis, MO).

Animals

Male C57BL/6 mice were provided by the Animal Center of Wenzhou Medical University, whereas GemPharmatech (Jiangsu, China) provided FGFR1 floxed mice on a C57BL6/JGpt background (FGFR1f/f; strain ID T009598) and mice with Lyz2-Cre knock-in at the H11 site on the same background (Strain ID T055107). Macrophage-specific FGFR1 knockout mice (FGFR1-CKO) were acquired by crossing FGFR1f/f and Lyz-Cre mice. All animal studies followed the review and approval of care and experimental procedures by the Wenzhou Medical University Animal Policy and Welfare Committee (Approval Document No. wydw2020-0117) and conformed to the NIH Guide for the Care and Use of Laboratory Animals.

Cell culture

Primary mouse peritoneal macrophages (MPMs) were isolated from FGFR1f/f and FGFR1-CKO mice, as described previously [17]. Briefly, these MPMs were cultured in RPMI-1640 augmented with penicillin (100 U/mL), streptomycin (100 mg/mL) (cat# 1514-0122, Invitrogen, Carlsbad, CA) and 10% fetal bovine serum (FBS, cat# 10099141 C, Gibco, Eggenstein, Germany) in a humidified incubator with 5% CO2 at 37 °C. Furthermore, human HSCs, LX-2, was acquired from Procell Life Science & Technology (Wuhan, China) and cultured in 4.5 g/L glucose DMEM supplemented with 20% FBS, penicillin (100 U/mL) and streptomycin (100 mg/mL) in 5% CO2 at 37 °C. Primary mouse hepatocytes were isolated from FGFR1f/f and FGFR1-CKO mice by a modified two-step collagenase perfusion procedure [18].

Development of NAFLD mouse model

For 24 weeks, a low-fat diet (LFD) or a high-fat diet (HFD) was given to 8-week-old male FGFR1-CKO and littermate FGFR1f/f male mice. Mice were then divided into four groups (n = 6/group): FGFR1-CKO-LFD, FGFR1-CKO-HFD, FGFR1f/f-LFD, and FGFR1f/f-HFD. The LFD group was fed a standard rodent diet containing 14.7 kcal.% protein, 9.4 fkcal.% at, and 75.9 kcal.% carbohydrates (cat #MD12031, MediScience Diets Co., Ltd., Yangzhou, China). The HFD comprised 60 kcal.% fat, 20 kcal.% protein, and 20 kcal.% carbohydrates (cat #MD12033, MediScience Diets Co., Ltd.).

For pharmacological analysis, the FGFR1 inhibitor, AZD4547, was administered in the NAFLD mice. The normal control C57BL/6 mice (Con, n = 6) received LFD fed for 24 weeks, whereas the HFD group (HFD, n  =  6) received HFD for 24 weeks. C57BL/6 mice in Con and HFD groups were treated with vehicle (1% CMC-Na) from the 16th to the 24th week. From 16th to 24th week, the HFD-fed mice were treated with AZD4547 (5  or 10 mg/kg, respectively) once every 2 days via gavage (HFD + AZD-5, HFD + AZD-10, n = 6/group). Furthermore, some LFD-fed mice (n = 6) were treated with AZD4547 (10 mg/kg) from the 16th to the 24th week, once every 2 days by gavage (AZD-10, n = 6). At week 24, the mice were sacrificed, and the blood and liver samples were obtained.

RNA-sequencing

Total RNA was isolated from liver tissues of FGFR1f/f-HFD, FGFR1f/f-LFD, and FGFR1-CKO-HFD mice using TRIzol reagent (cat# 15596018; Thermo Fisher; Waltham, MA). RNA-seq analysis was performed at Hangzhou Lc-Bio Technologies Co., Ltd, China.

Pathological analysis of liver tissues

Liver tissue samples were fixed in 4% paraformaldehyde, embedded in paraffin, sliced (5-μm thickness), dehydrated, and then stained with hematoxylin and eosin (H&E) to evaluate the general histopathological damage under light microscopy. Furthermore, some paraffin sections were also stained with Sirius Red and Masson’s Trichrome Stain to assess the levels of liver fibrosis.

Immunohistochemistry (IHC)

After the liver samples were rehydrated, they were subjected to antigen retrieval in 0.01 mol/L citrate buffer (pH 6.0) heated in microwave and then incubated for 30 min in 3% hydrogen peroxide in methanol at room temperature. Subsequently, the samples were blocked with 5% BSA (Sigma, Burbank, CA), treated with anti-F4/80 antibody (1:200) or anti-Ly-6G (1:200) at 4 °C overnight, washed, and then tagged with the corresponding secondary antibody (1:200) for 1 h. The reaction was visualized with DAB (ZSGB-Bio, Beijing, China) and counterstained with hematoxylin. The images were acquired using a microscope (Nikon, Tokyo, Japan).

Immunofluorescence staining

Frozen liver tissues in Optimal Cutting Temperature (OCT) media were sectioned at 5-μm thickness, fixed with formalin treatment for 10 min, blocked using 1% BSA for 1 h, and then incubated with primary antibodies at 4 °C overnight. Then, the sections were treated with fluorophore-conjugated secondary antibodies for 1 h at room temperature, counterstained with DAPI, and imaged using an epi-fluorescence microscope (Nikon, Tokyo, Japan).

Lipid staining in cells

First, the cells were fixed in 10% formalin for 10 min and then treated with 100% propylene glycol for 10 min at room temperature. After fixation, cells were stained with Oil Red O working solution for 10 min, and images were acquired using a microscope.

Cytokine measurements

To assess the effects of various treatments on cytokine secretion, in vitro studies were performed using the conditioned medium. Total protein in the collected cells and secreted TGF-β levels in MPMs culture media were quantified using the BCA Protein Assay (cat# 23225; Thermo Fisher, Waltham, MA) and a commercial TGF-β ELISA kit, respectively. Furthermore, in vivo, circulating cytokine levels were measured in serum samples using TNF-α and IL-6 ELISA kits. OD450 nm was measured using the SpectraMax M5 microplate reader (Molecular Devices; Silicon Valley, CA). The cytokine levels were normalized to the total protein measurements obtained.

Western blot

Nuclear extracts were acquired from MPMs using a cytoplasmic and nuclear protein isolation kit (cat# P0028; Beyotime Biological Technology, Shanghai, China). Cell and tissue lysates were prepared using RIPA buffer (cat# P0013B; Beyotime Biological Technology) with 1 × Halt Protease and Phosphatase Inhibitor Cocktail (Thermo Fisher, Waltham, MA). Then, the acquired lysate was centrifuged for 10 min at 12,000 rpm at 4 °C; the protein was quantified and subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) for separation. The separated proteins were then transferred to polyvinylidene fluoride membranes, which were blocked in Tris-buffered saline (pH 7.4, containing 0.05% Tween 20 and 5% non-fat milk) for 1.5 h at room temperature and incubated overnight at 4 °C with primary antibodies. Subsequently, the membranes were treated with horseradish peroxidase-conjugated secondary antibodies for 1 h at room temperature. Immunoreactivity was visualized using enhanced chemiluminescence reagent (Bio-Rad, Hercules, CA) and quantified using ImageJ software (v 1.53i, NIH, Bethesda, MD). Values were normalized to respective housekeeping proteins.

Real-time qPCR

Total cellular and tissue RNA was extracted using TRIzol Reagent (Thermo Fisher), reverse-transcribed using PrimeScript RT reagent (cat# RR047A, Takara Bioscience, Kusatsu, Japan) and then subjected to real-time PCR using TB Green Premix Ex Taq II (cat# RR820A, Takara) on the CFX96 Touch Real-Time PCR Detection System (Bio-Rad). The expression was normalized to Actb, and the relative expression levels were calculated using the 2−ΔΔCT method. Supplementary Table S1 provides a list of the primer sequences used.

MTT assay

The LX-2 cells were seeded in 96-well plates and cultured overnight at 37 °C. The cells were then treated with conditioned media from MPMs for varying periods. After 4 h of treatment, cell viability was measured using the MTT reagent (cat# M8180-250MG, Solarbio, Beijing, China); the reaction was stopped with N, N-dimethylformamide. The absorbance was measured at 490 nm to assess cell viability.

Statistical analysis

All statistical analyses were performed using GraphPad Prism 8.0 software (GraphPad, San Diego, CA), and the data are presented as mean ± SEM. Student’s t-test was used to compare two groups of data, whereas for more than two groups, one-way ANOVA followed by Dunnett’s post-hoc test was performed. P < 0.05 were considered to be statistically significant. Post-tests were run only if F achieved P < 0.05 and there was no significant variance inhomogeneity.

Results

High-fat-diet increases phosphorylation of FGFR1 in the mouse liver

To evaluate whether p-FGFR1 participates in NAFLD, the expression of p-FGFR1 in the liver tissues of NAFLD mice was analyzed. As Fig. 1a, b indicate, the expression of p-FGFR1 was markedly increased in the liver of the HFD-fed mice compared with LFD-fed mice. Consistently, immunoblotting showed that HFD increased the p-FGFR1 levels in liver tissues (Fig. 1c, d). Previous studies have reported that FGFR1 is expressed in macrophages [19–21]. Immunofluorescence staining data show that p-FGFR1 immunoreactivity was co-localized with F4/80 in the liver of HFD-fed C57BL/6 mice (Fig. 1e, Supplementary Fig. S1a); this was confirmed by repeating this experiment using livers from HFD-fed FGFR1-CKO mice. Although HFD increased macrophage infiltration in the liver, there was only a few p-FGFR1 expression in livers of HFD-fed FGFR1-CKO mice (Supplementary Fig. 1b). Additionally, the expression of p-FGFR1 did not co-localize with the expression of either HNF-4α (hepatocytes marker) or α-SMA (HSCs marker), indicating hepatocytes and HSCs are not the major cell types for p-FGFR1 expression (Supplementary Fig. S1c, d). Moreover, Western blot data confirmed that FGFR1 was primarily expressed in MPMs but not the primary hepatocytes or HSCs (Fig. 1f, Supplementary Fig. S1e). In addition, the protein levels of p-FGFR1 were only significantly increased after PA stimulation in MPMs (Fig. 1f, Supplementary Fig. S1e). All these data indicate that p-FGFR1 was mostly expressed in macrophages. Next, to confirm whether FGFR1 of macrophages is involved in the development of NAFLD, MPMs were treated with palmitate (PA), a common saturated fatty acid. Western blot results showed that PA induced FGFR1 phosphorylation in MPMs in a time-dependent manner (Fig. 1g, Supplementary Fig. S1f). These results indicate that HFD increased FGFR1 phosphorylation in liver macrophages, which may correlate with liver injury in NAFLD mice.

Fig. 1. Phosphorylation of FGFR1 is increased in macrophages in the liver of HFD-fed mice.

Fig. 1

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Liver tissues were harvested from C57BL/6 mice fed with a low-fat diet (LFD) or high-fat diet (HFD) for 24 weeks, respectively. a The expression of p-FGFR1 in the liver was examined using IHC [scale bar = 50 µm]. b The quantitative analysis of p-FGFR1 expression. c Representative Western blot images of p-FGFR1 and FGFR1 in liver proteins. GAPDH was used as the loading control. d Densitometric quantification of p-FGFR1. e Representative immunofluorescence staining images of liver tissues. Macrophage marker F4/80 is represented by green, and p-FGFR1 is indicated by red. Tissues were counterstained with DAPI (blue). [scale bar = 50 µm]. f Mouse primary macrophages (MPMs), hepatocytes, and hepatic stellate cells (HSCs) were treated with PA (200 μM) for 30 min. The protein levels of p-FGFR1 and FGFR1 were determined using Western blot assay. GAPDH was used as the loading control. g MPMs were treated with PA (200 μM) for 5, 15, 30, 60, and 120 min. Western blot assay of protein levels of p-FGFR1 and FGFR1 in MPMs. GAPDH was used as the loading control. Statistical data were presented as mean ± SEM; n = 3 or 6 for in vivo experiments; n = 3 for in vitro experiments; *P < 0.05.

FGFR1 knockout prevents HFD-induced liver lipid accumulation and fibrosis

In the development and progression of NAFLD, metabolic reprogramming in hepatic macrophages disrupts glycolysis, lipid synthesis, and iron metabolism, thereby causing liver steatosis, inflammation, and fibrogenesis [22–24]. To understand the role of FGFR1 in NAFLD, FGFR1-CKO and FGFR1f/f mice were either fed LFD or HFD for 24 weeks, and then their liver injury was examined. It was observed that HFD increased body weight in both FGFR1f/f and FGFR1-CKO mice (Supplementary Fig. S2a). Although macrophage-specific FGFR1 knockout did not reverse this body weight increase, it reduced the HFD-increased liver weight in FGFR1-CKO mice when compared with FGFR1f/f mice (Fig. 2a, c). Furthermore, the H&E staining revealed that fat accumulation was markedly reduced in the FGFR1-CKO mice after HFD treatment (Fig. 2b). According to the NAFLD activity score (NAS), macrophage-specific FGFR1 knockout alleviated HFD-induced injury, as indicated by lower NAS in HFD-treated FGFR1-CKO mice compared to HFD-treated FGFR1f/f mice (Supplementary Fig. S2b). HFD-treated FGFR1-CKO mice had significantly reduced levels of total cholesterol, LDL, and triglycerides than HFD-treated FGFR1f/f mice (Fig. 2d–f). In addition, serum ALT and AST levels were markedly reduced in HFD-fed FGFR1-CKO mice than those in HFD-fed FGFR1f/f mice (Fig. 2g, h). It has been suggested that liver fibrosis is an important phenotype of NAFLD, and it is markedly associated with FGFR1 [12, 25]. Therefore, whether FGFR1 affected liver fibrosis was evaluated in vivo. Both Sirus Red and Masson’s Trichrome staining results indicated that HFD triggered fibrosis only in FGFR1f/f mice, while FGFR1-CKO significantly reduced HFD-induced hepatic fibrosis (Fig. 2i-j, Supplementary Fig. S2c, d). Consistent with these structural alterations, HFD-fed FGFR1-CKO mice indicated significantly alleviated protein and mRNA levels of fibrosis-associated factors, including CoL-1, α-SMA, and TGF-β, suggesting the protective role of macrophage-specific FGFR1 knockout in NAFLD mice (Fig. 2k, l, Supplementary Fig. S2e). Similarly, in HFD-fed FGFR1-CKO mice, the serum TGF-β levels were reduced compared with the FGFR1f/f mice during HFD-induced fibrosis progression (Supplementary Fig. S2f).

Fig. 2. Macrophage-specific FGFR1 knockout protects against HFD-induced fibrosis and lipid accumulation in liver.

Fig. 2

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FGFR1-CKO and FGFR1f/f mice were fed LFD or HFD for 24 weeks. a Representative images of mouse liver tissues. b Liver sections were stained with H&E [scale bar = 50 µm]. c The ratio of liver weight to tibia length (LW/TL; mg/mm). d–f Serum levels of total cholesterol (TCH), low-density lipoprotein-cholesterol (LDL-C), and triglyceride (TG) in mice. g, h Serum levels of liver function markers ALT and AST in mice. i, j Levels of liver fibrosis were determined using Sirius Red and Masson staining [scale bar = 50 µm]. k Immunoblotting shows levels of fibrosis-associated factors, including α-SMA, TGF-β, and COL-1, in the liver. GAPDH was used as the loading control. l mRNA levels of fibrosis-associated genes such as Col1a1, Tgfb1, and Acta2 in the liver were measured using RT-qPCR. Transcripts were normalized to that of Actb. Statistical data were presented as mean ± SEM; n = 6; *P < 0.05.

Macrophage-specific FGFR1 knockout protects against HFD-induced liver inflammation by suppressing the activation of MAPKs and TNF signaling pathways

To investigate the potential mechanism underlying the protective effect of FGFR1-CKO against HFD-induced liver injury, RNA-sequencing analysis of NFALD tissues was performed. A total of 1489 differentially expressed genes (DEGs), including 628 upregulated and 861 downregulated genes, were identified when FGFR1f/f-HFD and FGFR1f/f-LFD groups were compared (Supplementary Fig. S2g). Consistently, 1006 DEGs (573 upregulated and 433 downregulated) were identified after FGFR1-CKO-HFD and FGFR1f/f-HFD comparison (Supplementary Fig. S2h). Interestingly, KEGG enrichment analysis revealed that the MAPKs and TNF signaling pathways were common in both the FGFR1f/f-LFD to FGFR1f/f-HFD and FGFR1-CKO-LFD to FGFR1-CKO-HFD groups (Fig. 3a, Supplementary Fig. 2i), indicating the involvement of these signaling pathways in FGFR1-mediated liver protection. Furthermore, it was found that serum TNF-α and IL-6 levels (Fig. 3b) and the mRNA levels of Tnf, Il6, and Il1b were increased in HFD-fed FGFR1f/f mice but not in HFD-fed FGFR1-CKO mice (Fig. 3c–e). IHC data revealed that F4/80 immunoreactivity was decreased in the liver of HFD-fed FGFR1-CKO mice (Fig. 3f, Supplementary Fig. S3a). Moreover, the macrophage infiltration data also indicated decreased Ly-6G in HFD-fed FGFR1-CKO mice, suggesting macrophage-specific FGFR1 knockout also inhibited the neutrophil infiltration (Fig. 3g, Supplementary Fig. S3b). Furthermore, Western blotting data validated the increased phosphorylation of ERK and JNK in HFD-fed FGFR1f/f mice, while FGFR1-CKO significantly inhibited the phosphorylation of MAPKs in the liver (Fig. 3h, Supplementary Fig. S3d). The literature has indicated that NF-κB is a key component of the TNF signaling pathways [26]. Therefore, the levels of phosphorylated P65 and IκBα degradation in the liver were also evaluated, which indicated reduced p-P65 and increased IκB-α levels in liver lysates of HFD-fed FGFR1-CKO mice (Fig. 3i, Supplementary Fig. S3e).

Fig. 3. Macrophage-specific FGFR1 knockout prevents HFD-induced liver inflammation by suppressing MAPK and TNF signaling pathways.

Fig. 3

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a Venn diagram of KEGG analysis of FGFR1f/f-HFD group vs. FGFR1f/f-LFD group and FGFR1-CKO-HFD group vs. FGFR1-CKO-LFD group. b Serum levels of TNF-α and IL-6 in FGFR1f/f and FGFR1-CKO mice fed with LFD or HFD. c–e mRNA levels of Tnf, Il6 and Il1b in liver. f, g Representative images of liver sections stained for macrophage marker F4/80 (brown) and Ly-6G (brown). Sections were counterstained with hematoxylin (blue) [scale bar = 50 μm]. h Immunoblot analysis of p-JNK, JNK, p-ERK, and ERK in liver proteins. GAPDH was used as the loading control. i Liver lysates were probed for p-P65, P65, and IκB-α. GAPDH was used as the loading control. Statistical data were presented as mean ± SEM; n = 6; *P < 0.05.

FGFR1 deficiency inhibits PA-induced inflammatory responses in macrophages

The HFD-induced activation of macrophages increases inflammatory cytokine production [27]. Thus, the anti-inflammatory effect of FGFR1 in PA-treated MPMs from FGFR1f/f and FGFR1-CKO mice were further assessed. It was revealed that FGFR1 deficiency reduced the protein levels of TNF-α and IL-6 in MPMs from FGFR1-CKO mice (Fig. 4a). Furthermore, PA-induced mRNA levels of inflammatory cytokines, adhesion molecules, and chemokines were significantly lower in MPMs derived from FGFR1-CKO than in those derived from FGFR1f/f mice (Fig. 4b, c). Western blot results showed that FGFR1 knockout in MPMs significantly inhibited the activation of MAPKs and TNF signaling pathways (Fig. 4d, Supplementary Fig. S3f, g). Moreover, it was revealed that PA stimulation markedly promoted P65 nuclear translocation in MPMs derived from FGFR1f/f mice. However, FGFR1-CKO mice-derived MPMs notably suppressed the PA-induced nuclear translocation of P65 (Fig. 4e, f). These results show that PA-induced FGFR1-mediated downstream activation of MAPKs and TNF pathways leads to inflammatory factor expression in both infiltrating and resident macrophages.

Fig. 4. FGFR1 deficiency suppresses PA-induced inflammatory responses in macrophages.

Fig. 4

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MPMs were isolated from FGFR1f/f and FGFR1-CKO mice. a MPMs were exposed to 200 μM PA or 10% BSA (CON) for 24 h. Protein levels of TNF-α and IL-6 in the culture medium were determined via ELISA. b, c RT-qPCR analysis of mRNA levels of inflammatory factors and adhesion molecules after MPMs were exposed to 200 μM PA or 10% BSA (CON) for 12 h. Transcripts were normalized to Actb. d Protein levels of p-JNK, JNK, p-ERK, ERK, p-P65, P65, and IκB-α were analyzed using Western blot assay after MPMs were exposed to 200 μM PA or 10% BSA for 30 or 60 min. GAPDH was used as the loading control. e MPMs were exposed to 200 μM PA or 10% BSA for 30 min. Cytoplasmic and nuclear proteins were collected to measure the expression of p65. GAPDH and Lamin B were used as the loading controls. f Densitometric quantification for the blots in (e). Statistical data were presented as mean ± SEM; n = 3; *P < 0.05.

Macrophage-specific FGFR1 knockout reduces fat deposition and inflammatory responses in hepatocytes

It has been indicated that macrophage-induced cytokines release can directly affect hepatocytes and promote steatosis, inflammation, and hepatocellular damage [28]. Therefore, how macrophages induce liver parenchymal cell change after PA stimulation were further studied. MPMs from FGFR1f/f and FGFR1-CKO mice were cultured and treated with PA. The CM of PA-stimulated macrophages was acquired and added in C57BL/6 mice-derived primary mouse hepatocytes. The hepatocytes were then collected for further evaluation. The schematic graphic illustration of this experiment is shown in Fig. 5a. Oil Red O analysis indicated lipid accumulation in hepatocytes treated with CM from FGFR1f/f MPMs, which was significantly reduced in hepatocytes treated with CM of FGFR1-CKO MPMs (Fig. 5b). Furthermore, hepatocytes exposed to CM of FGFR1-CKO MPMs also indicated marked reduction in the induction of lipid metabolism genes, including Acaca, Srebp1, Pparα, and Cpt1α (Fig. 5c). Moreover, it was found that MAPKs and TNF pathways were inhibited under exposure to CM from FGFR1-CKO MPMs (Fig. 5d, Supplementary Fig. S4c, d). Additionally, mRNA levels of inflammatory cytokines and chemokines were greatly decreased in hepatocytes upon exposure to CM from FGFR1-CKO MPMs (Fig. 5e, f). It is reported that lipid metabolism genes in hepatocytes are often regulated by AMPK [29]. Therefore, the primary liver cells were challenged with CM from PA-stimulated macrophages to determine the activation of the AMPK pathway. It was observed that AMPK phosphorylation was significantly reduced in hepatocytes treated with CM of PA-induced FGFR1f/f MPMs but increased in those treated with CM of PA-induced FGFR1-CKO MPMs (Supplementary Fig. S4a, b). These data suggest that macrophage-specific FGFR1 knockout also plays a role in the regulation of the AMPK pathway. Altogether, the data suggest that FGFR1 is essentially associated with hepatocyte lipid metabolic dysfunction and inflammatory response in macrophages.

Fig. 5. Factors released from PA-stimulated macrophages cause lipid accumulation and inflammatory responses in hepatocytes via an FGFR1-dependent manner.

Fig. 5

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a Schematic showing experimental design. MPMs derived from FGFR1f/f and FGFR1-CKO mice were exposed to 200 μM PA or vehicle control for 24 h. Conditioned medium (CM) was collected and applied to primary hepatocytes at a 1:1 ratio with fresh medium. Hepatocytes were examined after 24 h stimulation. b Accumulated lipid in hepatocytes was determined via Oil Red O staining [scale bar = 50 µm]. c mRNA levels of genes regulating fat deposition. Transcripts were normalized to that of Actb. d Protein levels of p-JNK, JNK, p-ERK, ERK, p-P65, P65, and IκB-α in hepatocytes were determined after CM was collected and added to primary hepatocytes at a 1:1 ratio with fresh medium for 60 min. GAPDH was used as the loading control. e, f mRNA levels of inflammatory factors in hepatocytes were measured after CM was collected and applied to primary hepatocytes at a 1:1 ratio with fresh medium for 24 h. Transcripts were normalized to that of Actb. Statistical data were presented as mean ± SEM; n = 3; *P < 0.05.

Macrophage-specific FGFR1 knockout suppresses activation of hepatic stellate cells

The HSCs are widely recognized as key contributors to hepatic fibrogenesis [30, 31], and TGF-β is a major factor that activates HSCs [32, 33]. Therefore, the levels of TGF-β in CM of FGFR1f/f and FGFR1-CKO MPMs exposed to PA were measured. As shown in Fig. 6a, TGF-β production was decreased in the culture supernatant of macrophages from FGFR1-CKO mice compared with those from FGFR1f/f mice, suggesting that in the absence of FGFR1, macrophages may affect the activation of stellate cells. To test this hypothesis, the same CM conditional culture system to HSC cell line, LX-2 cells, was utilized. The schematic graphic illustration of this experiment is shown in Fig. 6b. It was indicated that CM generated from PA-treated FGFR1f/f MPMs increased the proliferation of LX-2 cells, while CM from PA-treated FGFR1-CKO MPMs did not (Fig. 6c). Protein markers of HSCs activation and fibrosis increased after LX-2 cells were exposed to FGFR1f/f MPMs CM, but not after exposure to FGFR1-CKO MPMs CM (Fig. 6d, Supplementary Fig. S5a). Furthermore, mRNA levels of fibrosis-associated factors also showed similar results (Fig. 6e–g).

Fig. 6. FGFR1 is required for macrophages to activate hepatic stellate cells.

Fig. 6

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a MPMs harvested from FGFR1f/f and FGFR1-CKO mice were exposed to 200 μM PA or 10% BSA (CON) for 24 h. TGF-β levels were determined in culture medium via ELISA [n = 4]. b Schematic showing the experimental setup to assess hepatic stellate cell (HSCs) activation. MPMs harvested from FGFR1f/f and FGFR1-CKO mice were exposed to 200 μM PA or vehicle control for 24 h. CM was collected and added to the LX-2 cells at a 1:1 ratio with fresh medium, which was then collected for further evaluation. c Proliferation of LX-2 cells was measured at different time points [n = 3]. d Lysates prepared from LX-2 cells were used to examine the protein levels of α-SMA, COL-1, and TGF-β via immunoblotting. GAPDH was used as the loading control. e-g mRNA levels of fibrosis-associated genes in LX-2 cells were measured via RT-qPCR. Transcripts were normalized to that of Actb [n = 3]. Statistical data were presented as mean ± SEM; *P < 0.05.

FGFR1 inhibitor alleviates HFD-induced liver injury by reducing lipid accumulation, fibrosis and inflammation

AZD4547 is an orally bioavailable selective inhibitor of FGFR tyrosine kinases [16]. The pharmacological inhibition of FGFR1 was assessed as an effective treatment option for HFD-induced liver injury. H&E, Sirius Red, and Masson’s Trichrome staining analysis of the liver showed that AZD4547 reduced lipid accumulation and fibrosis in the HFD-fed mice (Fig. 7a–e). Furthermore, NAS was significantly reduced after AZD4547 administration in HFD-fed WT mice (Supplementary Fig. S5c). In addition, AZD4547 treatment reduced the HFD-induced liver weight increase in a dose-dependent manner (Fig. 7f). Moreover, AZD4547 administration significantly lowered the serum ALT and AST levels compared with the HFD group, indicating preservation of liver function after FGFR1 blockade (Fig. 7g, h). The serum lipid profile showed a beneficial effect of AZD4547 in HFD-fed mice (Fig. 7i–k). Additionally, protein and mRNA levels of fibrosis-associated factors (Col-1, α-SMA, and TGF-β) confirmed the protective role of AZD4547 treatment against NAFLD (Fig. 7l, m, Supplementary Fig. S5d). The immunostaining of F4/80 and Ly-6G indicated decreased macrophage infiltration in liver of AZD4547-treated mice (Fig. 8a–c, Supplementary Fig. S5e). Additionally, protein and mRNA levels of inflammatory cytokines were significantly reduced after AZD4547 treatment (Fig. 8d–h, Supplementary Fig. S5f). Lastly, the pharmacological mechanism of AZD4547 was also similar to that identified in FGFR1-CKO mice. AZD4547 treatment suppressed HFD-induced activation of MAPKs and TNF pathways in the liver (Fig. 8i, j, Supplementary Fig. S5g, h). Altogether, the data validated that the pharmacological inhibition of FGFR1 by AZD4547 had a therapeutic role in HFD-induced liver injury by inhibiting lipid accumulation, inflammation, and fibrosis.

Fig. 7. FGFR1 inhibitor alleviates HFD-induced liver injury.

Fig. 7

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C57BL/6 mice were fed with LFD (CON) or HFD for 24 weeks with or without the treatment of AZD4547 starting at week 16 (HFD + AZD-5 mg/kg, HFD + AZD-10 mg/kg). CMC-Na vehicle was administered in CON and HFD mice as controls. H&E (a), Sirius Red (b) and Masson (c) staining of liver sections were shown [scale bar = 50 µm]. Quantification of Sirius Red (d) and Masson staining (e) is shown. f Measurement of liver weight to tibia length (LW/TL; mg/mm). g, h Serum levels of liver function markers such as ALT (g), AST (h), TG (i), TCH (j) and LDL-C (k) in mice. l, m Protein and mRNA levels of fibrosis-associated factors in liver tissues. Statistical data were presented as mean ± SEM; n = 6; *P < 0.05.

Fig. 8. FGFR1 inhibitor prevents HFD-induced liver inflammation through inhibiting the activation of MAPK and TNF pathways.

Fig. 8

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The mice were treated as described in Fig. 7. a, b Representative liver sections stained for macrophage marker F4/80 (brown) and Ly-6G (brown). Sections were counterstained with hematoxylin (blue) [scale bar = 50 µm]. c Quantification of staining of F4/80 is shown. d, e Serum levels of TNF-α and IL-6 were measured via ELISA. f–h mRNA levels of inflammatory cytokines in liver. i Immunoblot analysis of p-JNK and p-ERK in liver. ERK, JNK and GAPDH were used as the loading controls. j Liver lysates were probed for p-P65 and IκB-α. P65 and GAPDH were used as loading controls. Statistical data were presented as mean ± SEM; n = 6; *P < 0.05.

Discussion

This study examined the potential role of FGFR1 in the development of NAFLD and indicated that FGFR1 expression was elevated in the liver of HFD-induced NAFLD mice. Specifically, targeting FGFR1 in macrophages alleviated HFD-induced inflammation, lipid accumulation, and fibrosis in the liver, improving liver function and performance. Furthermore, FGFR1 inhibition in macrophages produced these therapeutic benefits by inactivation of MAPKs and TNF pathways, thereby reducing the production of pro-inflammatory cytokines and TGF-β expression in macrophages. The FGFR1-mediated release of cytokines stimulated the accumulation of lipids in hepatocytes, while macrophage-secreted TGF-β triggered the activation of HSCs, ultimately resulting in liver fibrosis. Furthermore, the FGFR1 inhibitor indicated a significant protective effect on the liver in NAFLD mice. These data suggest that targeting FGFR1 may be a promising therapeutic strategy for treating NAFLD.

It has been indicated that obesity causes chronic low-grade inflammation, particularly in adipose tissue and liver [34], and macrophages are the dominant immune cell type causing inflammation in obese and T2DM islets [35]. Resident and recruited macrophages in liver are associated with the pathogenesis of various liver diseases [36]. Bone marrow-derived macrophages increase in number during liver injury [37]. Therefore, inhibiting macrophage-mediated inflammation is an effective strategy for treating obesity-related liver diseases. This research indicated that phosphorylated FGFR1 was strongly expressed in F4/80 positive cells in the liver of NAFLD mice. Specific knockdown of FGFR1 in macrophages demonstrated that macrophage-specific FGFR1 is associated with lipid metabolism, inflammation and fibrosis in the liver of NAFLD mice. Mechanistically, FGFR1 affected HFD-induced macrophage activation and increased inflammatory cytokines levels. At the cellular level, MPMs released pro-inflammatory cytokines exposed to PA were reduced by inhibiting FGFR1. Another research also demonstrated that macrophage-specific deletion or pharmacological inhibition of MyD88 prevents liver damage in NAFLD by reducing inflammatory response [38]. This research indicates that macrophage-mediated inflammatory responses are critical in NAFLD, consistent with this present study, which suggests that FGFR1 significantly contributed to the pathogenesis of NAFLD via macrophage-induced inflammation in liver.

To investigate the potential mechanisms underlying the liver-protective effects of FGFR1, RNA-sequencing analysis was performed, which revealed that the MAPKs and TNF signaling pathways might intersect. The literature indicates that PA regulates the inflammatory response by activating MAPKs and NF-κB signaling pathways [39], the activation of which promotes liver fibrosis and lipid accumulation, exacerbating liver damage [40]. Several studies have shown that regulation of MAPKs and NF-κB signaling pathways improves liver damage, inflammation, fibrosis, and steatosis caused by obesity-related metabolic disorders [41–44]. Wang et al. revealed that FGFR inhibition suppresses TNFα-induced activation of the NF-κB pathway in HSCs and inhibits inflammation due to acute liver injuries [12]. FGFR1-mediated MAPK activation causes cholestatic injury in liver sinusoidal endothelial cells (LSECs), resulting in CXCR4-dominated pro-fibrotic transition of angiocrine response during liver repair [25]. These studies highlight that FGFR1 in macrophages might serve as a potential target for NAFLD treatment. Here, it was indicated that HFD increased the activation of phosphorylated MAPKs (ERK, JNK) and NF-κB in the liver, MPMs, and cultured primary hepatocytes, which could be significantly inhibited by macrophage-specific deletion of FGFR1. Furthermore, lipid accumulation and liver fibrosis were significantly alleviated.

AZD4547 is a well-known selective small molecule inhibitor of FGFR1, which is significantly effective against FGFR-dysregulated tumors in preclinical models [16]. Besides, it has been reported to exhibit many other pharmacological effects as well. For example, AZD4547 is shown to exert a protective effect against excessive inflammatory damage in septic mice by inhibiting the LPS-induced phosphorylation of key NF-κB/MAPK/STAT3 pathways [45]. AZD4547 also attenuates lipopolysaccharide-induced acute kidney injury by inhibiting inflammation [46]. Our previous study indicated that upon the Ang II challenge, AZD4547 protects kidneys from dysfunction and fibrosis [47]. Here, it was revealed that AZD4547 is equally effective in reducing NAFLD manifestations as macrophage-specific FGFR1 knockout without significant toxic side effects. Since it has been proved that p-FGFR1 was mainly found in macrophages, we believe that the oral AZD4547 administration mostly inhibited the phosphorylation of FGFR1 in macrophages but not in the hepatocytes or HSCs. Overall, this is the first study to reveal that AZD4547 can be used to treat NAFLD, which is a new finding in this field.

Some limitations of this research are: firstly, how PA acts on FGFR1 to cause its phosphorylation was not clear. FGFR1 is a transmembrane protein that contains an extracellular ligand-binding domain, a single transmembrane helix, and a cytosolic region with tyrosine kinase activity [8]. Therefore, it is worth exploring whether PA directly binds to FGFR1. Secondly, how the non-classic ligand, PA, induces FGFR1-mediated inflammatory response in the present study was also not assessed. After being activated by the classic FGF ligand family, FGFRs phosphorylate specific tyrosine residues that mediate interaction with cytosolic adaptor proteins and the RAS-MAPK, PI3K-AKT, PLCγ, and STAT intracellular signaling pathways [48]. The RNA sequencing analysis indicated that MAPKs and TNF signaling pathways regulate FGFR1 to exert its anti-inflammatory effects in NAFLD. Thus, future studies are needed to clarify the relationship between the classic FGFR1 signaling and MAPKs/TNF signaling pathways.

Conclusion

In summary, this study provides a comprehensive understanding of FGFR1’s role in NAFLD. Through genetic knockout or pharmacological intervention of FGFR1 in macrophages, it was demonstrated that FGFR1 plays a critical role in HFD-induced inflammatory responses in macrophages. Furthermore, it was found that after FGFR1 blockade in macrophages, lipid accumulation in hepatocytes, and associated fibro-inflammatory responses in the liver are significantly reduced. These findings provide significant evidence of the importance of FGFR1 in the pathology of NAFLD and suggest that targeting FGFR1 could be a potential clinical therapeutic strategy.

Supplementary information

Supplementary file (10.1MB, zip)

Acknowledgements

Financial support was provided by the Medical and Health Science Research Project of Zhejiang Province (2023XY164 to LJH, 2022KY348 to WZ), Key Research Project of Wenzhou City (ZY2021021 to YW).

Author contributions

YNZ, ZDL, TY, and YW contributed to the literature search and study design. YNZ, TXX, TYJ, YSJ, WZ, KYL, and YW performed the experiments and analyzed the data. LJH and KYL provided technical help. TXX, TYJ, LJH, and YW participated in the drafting of the article. All authors agree to be accountable for all aspects of work ensuring integrity and accuracy.

Competing interests

The authors declare no competing interests.

Contributor Information

Kwang Youl Lee, Email: kwanglee@chonnam.ac.kr.

Li-jiang Huang, Email: 13777030956@163.com.

Yi Wang, Email: yi.wang1122@wmu.edu.cn.

Supplementary information

The online version contains supplementary material available at 10.1038/s41401-024-01226-7.

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