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Article

Human FGF1ΔHBS Gene Therapy as Treatment for Metabolic Dysfunction-Associated Steatohepatitis in ApoE-KO Mice

by
Yingjian Li
1,†,
Xiaodan Hui
1,†,
Chunjie Gu
1,
Qian Lin
2,
Ahmed Abdelbaset-Ismail
1,
Zixuan Xu
1,
Suchen Yadav
1,
Hongbiao Huang
1,
Jason Xu
1,
Sara E. Watson
3,4,
Kupper A. Wintergerst
3,4,5,
Lu Cai
1,3,5,6,7,
Zhongbin Deng
8,9,* and
Yi Tan
1,3,6,*
1
Pediatric Research Institute, Department of Pediatrics, School of Medicine, University of Louisville, Louisville, KY 40202, USA
2
Touchstone Diabetes Center, The University of Texas Southwestern Medical Center at Dallas, Dallas, TX 75390, USA
3
Wendy Novak Diabetes Institute, Norton Children’s Hospital, Louisville, KY 40202, USA
4
Norton Children’s Endocrinology, Department of Pediatrics, University of Louisville, Norton Children’s Hospital, Louisville, KY 40202, USA
5
The Center for Integrative Environmental Health Sciences, School of Medicine, University of Louisville, Louisville, KY 40202, USA
6
Department of Pharmacology and Toxicology, School of Medicine, University of Louisville, Louisville, KY 40202, USA
7
Department of Radiation Oncology, School of Medicine, University of Louisville, Louisville, KY 40202, USA
8
Division of Immunotherapy, Department of Surgery, School of Medicine, University of Louisville, Louisville, KY 40202, USA
9
Brown Cancer Center, School of Medicine, University of Louisville, Louisville, KY 40202, USA
*
Authors to whom correspondence should be addressed.
†
These authors contributed equally to this work.
Cells 2026, 15(5), 387; https://doi.org/10.3390/cells15050387
Submission received: 30 December 2025 / Revised: 8 February 2026 / Accepted: 17 February 2026 / Published: 24 February 2026
Browse Figures
Versions Notes

Abstract

The prevalence of metabolic dysfunction-associated steatohepatitis (MASH) is rising worldwide. hFGF1ΔHBS, a variant of human fibroblast growth factor 1 with three substitutions in its heparin-binding sites, was previously shown by our group to ameliorate fatty liver. However, hFGF1ΔHBS also significantly modulates systemic metabolism, making it unclear whether its hepatic benefits arise from direct liver-specific actions. Additionally, its poor pharmacokinetic profile underscores the need for alternative delivery strategies. Here, we employed adeno-associated virus serotype 8 under the thyroxine-binding globulin promoter (AAV8-TBG) to achieve sustained, hepatocyte-specific expression of hFGF1ΔHBS. In high-fat-, high-cholesterol-diet-fed apolipoprotein E knockout mice, liver-directed hFGF1ΔHBS expression markedly reduced hepatic steatosis, inflammation, and fibrosis, independent of changes in body weight, blood glucose, insulin sensitivity, body composition, or circulating triglyceride and cholesterol levels. Mechanistically, hFGF1ΔHBS gene transfer normalized fatty acid synthesis and suppressed fatty acid uptake by downregulation of stearoyl-CoA desaturase-1 and cluster of differentiation 36. Importantly, these therapeutic effects were achieved without inducing hepatic hyperproliferation, as evidenced by unchanged expression of proliferating cell nuclear antigen and antigen Kiel 67. Collectively, our findings demonstrate that hFGF1ΔHBS exerts direct hepatoprotective effects and that AAV8-TBG-mediated liver-directed hFGF1ΔHBS delivery represents a safe and effective strategy for treating MASH.

1. Introduction

The prevalence of metabolic dysfunction-associated steatohepatitis (MASH) is growing at an alarming rate, and MASH has become a major health problem worldwide [1]. MASH, characterized by hepatic lipid accumulation, inflammation, and fibrosis, is closely linked to systemic metabolic disturbances, including impaired glucose tolerance, insulin resistance and dysregulated lipid metabolism [2,3,4].
Fibroblast growth factor (FGF) 1 [5], a member of the paracrine FGF subfamily [6], has gained attention as a potential therapeutic molecule due to its documented roles in ischemic repair [7,8], wound healing [9,10], and neuroprotection [11]. Despite this promise, FGF1 remains preclinical or early clinical development, with no approved systemic therapies to date. In China, a topical formulation of FGF-1 has been approved for treating second-degree burns and chronic skin ulcers, where it promotes wound repair and tissue regeneration [12]. More recently, FGF1 has garnered increasing attention for its unexpected metabolic functions [13], including the regulation of blood glucose, enhancement of insulin sensitivity, and remodeling of adipose tissue [14,15]. These findings highlight its therapeutic potential for treating metabolic disorders such as obesity, diabetes and diabetic complications [16,17,18,19]. However, native FGF1 exhibits robust mitogenic activity, raising concerns about potential tumorigenic risk during long-term treatment. To address this limitation, we previously engineered a non-mitogenic variant, FGF1ΔHBS, which contains three substitutions within the heparin-binding site [20]. This variant markedly attenuates mitogenic activity while fully preserving the metabolic functions of wild-type FGF1 [20]. Notably, our recent studies showed that recombinant FGF1ΔHBS significantly improves nonalcoholic fatty liver disease (NAFLD) [21]. Nevertheless, because FGF1ΔHBS exerts potent systemic metabolic effects, it remains unclear whether its hepatic benefits are mediated directly through liver-intrinsic actions in vivo.
Another challenge limiting the therapeutic application of FGF1 is its short circulating half-life. Although direct pharmacokinetic data for recombinant FGF1ΔHBS in vivo remains limited, studies using an FGF1ΔHBS-containing chimeric molecule (FGF1ΔHBS–FGF21C-tail) report a circulating half-life of approximately 1.99 h [22]. Consequently, while FGF1ΔHBS represents a safer alternative to wild-type FGF1 [20], its rapid clearance necessitates long-term, repeated dosing, potentially reducing patient adherence and increasing the risk of immunogenicity from repeated exposure to exogenous proteins.
Adeno-associated virus serotype 8 (AAV8) vectors, particularly those incorporating the thyroxine-binding globulin (TBG) promoter, are widely used for liver-targeted gene delivery owing to their high hepatotropism and robust hepatocyte-restricted transcriptional activity [23,24,25]. The AAV8 capsid enables highly efficient transduction of murine hepatocytes following systemic administration, while the TBG promoter ensures selective transgene expression with minimal off-target activity in non-hepatic tissues [23,24,25]. Together, these features allow AAV8-TBG vectors to achieve sustained and potent hepatocyte-specific transgene expression that can persist for months after a single intravenous dose [23,24,25].
In this study, we leveraged AAV8 vectors carrying the TBG promoter to achieve efficient, liver-directed, long-term expression of the FGF1ΔHBS variant as a gene therapy strategy for MASH. This approach overcomes the pharmacokinetic limitations of recombinant protein delivery and enables direct evaluation of the hepatic actions and therapeutic potential of FGF1ΔHBS in the context of MASH.

2. Materials and Methods

2.1. Animals

Ten-week-old male ApoE-KO mice purchased from The Jackson Laboratory (Bar Harbor, ME, USA) were used to establish the MASH mouse model. Animals were maintained at 22 ± 2 °C under a 12 h light/dark cycle. Mice were randomly divided into two groups: MASH (n = 19), which was induced by feeding a high-fat, high-cholesterol (HFHC) diet (Catalog No. TD88137, Envigo, Indianapolis, IN, USA), and control mice (n = 6), which were maintained on a standard chow diet (Catalog No. TD.08485, Envigo, Indianapolis, IN, USA). Body weight and blood glucose levels were measured at the indicated time points, with blood glucose assessed using a FreeStyle complete blood glucose monitor (Abbott Diabetes Care Inc., Alameda, CA, USA). For tissue collection, mice were anesthetized, and the relevant tissues were excised and either snap-frozen at −80 °C or fixed in formalin for further analysis. All experimental procedures were approved by the Institutional Animal Care and Use Committee of the University of Louisville.

2.2. Recombinant AAV Vectors and Administration of AAV Vectors

Recombinant AAV8 vectors were generated by OBiO Technology (Sugar Land, TX, USA). Two constructs were used: AAV8-TBG-hFGF1ΔHBS-3 × FLAG-P2A-ZsGreen (AAV8-hFGF1ΔHBS, Plasmid ID: H42848; titer 1.84 × 1013 vg/mL), which expresses the non-mitogenic hFGF1ΔHBS variant containing three heparin-binding site substitutions, as previously described [20], and AAV8-TBG-P2A-ZsGreen (AAV8-Control, Plasmid ID: GL3009; titer 5.19 × 1013 vg/mL), which served as the control vector. All vectors were packaged in the AAV8 serotype and driven by the hepatocyte-specific TBG promoter. For systemic administration, AAV vectors were diluted in 200 μL of sterile PBS and administered via the tail-vein injection after 4 weeks of HFHC diet or chow feeding. MASH mice received either 4 × 109 vg/mouse (low dose, n = 6) or 2 × 1010 vg/mouse (high dose, n = 6) of AAV8-hFGF1ΔHBS, or 2 × 1010 vg/mouse of the AAV8-Control vector (n = 7). Control mice received AAV8-Control vector (n = 6). Following injection, mice remained on their assigned diets for an additional 10 weeks before euthanasia and tissue collection.

2.3. Intraperitoneal Glucose Tolerance Test (IPGTT), Intraperitoneal Insulin Tolerance Test (IPITT), and Dual-Energy X-Ray Absorptiometry (DEXA) Body Composition Analysis

Mice were fasted for 6 h prior to metabolic testing. For IPGTT and IPITT, animals received an intraperitoneal injection of glucose (1 g/kg body weight) or insulin (1 U/kg body weight). Blood glucose levels were measured at 0, 15, 30, 60, and 120 min after injection. Body composition was assessed using DEXA in whole-body scan mode, and lean and fat mass were quantified using the manufacturer’s software following standardized positioning protocols.

2.4. Biochemical Analysis for Plasma and Liver Samples

Plasma alanine aminotransferase (ALT) and aspartate aminotransferase (AST) levels were measured using commercially available kits (Cayman Chemical, Ann Arbor, MI, USA) according to the manufacturer’s instructions. Plasma samples were diluted at 1:2 in Assay Buffer (1×). For each reaction, 150 µL of ALT or AST substrate, 20 µL of ALT or AST cofactor, and 20 µL of sample were added to each well of a 96-well plate. After incubation at 37 °C for 15 min, 20 µL of ALT or AST Initiator was added, and the absorbance at 340 nm was recorded every minute for 10 min at 37 °C. Plasma and hepatic triglyceride levels were quantified using commercially available colorimetric kits (Cayman Chemical, Ann Arbor, MI, USA). Plasma samples were diluted 1:3 in the triglyceride reagents. For hepatic triglycerides, liver tissue (80–100 mg) was homogenized in 500 µL diluted NP40 reagent. For each reaction, 150 µL of the reconstituted and diluted triglyceride enzyme mixture and 10 µL of sample were added to a 96-well plate. After incubation at room temperature for 60 min, absorbance at 545 nm was measured using a microplate reader. Hepatic cholesterol content was measured using a colorimetric kit (Bio-Techne, Minneapolis, MN, USA). Liver tissue (20 mg) was homogenized in 180 µL anhydrous ethanol. A total of 250 µL of enzyme working solution and 2.5 µL of sample were added to each well of a 96-well plate. Following incubation at 37 °C for 10 min, absorbance at 510 nm was measured using a microplate reader. Plasma cholesterol levels were quantified using a colorimetric kit (Cell Biolabs, Inc., San Diego, CA, USA). Plasma samples were diluted at 1:50 in Assay Buffer, mixed, and incubated for 45 min at 37 °C. Fluorescence was measured immediately using a microplate reader with excitation at 530–570 nm and emission at 590–600 nm.

2.5. Liver Histopathological Analysis

Liver tissues were either fixed in 10% formalin for 48 h and embedded in paraffin or embedded in OCT and stored at −80 °C. For paraffin-embedded samples, sections were deparaffinized, rehydrated, and subjected to hematoxylin and eosin (H&E), Sirius Red, or Masson’s trichrome staining. Positively stained areas were quantified using Image J version 1.54p (NIH, Bethesda, MD, USA) and normalized to values obtained from control mice. For frozen sections, tissues were fixed in formalin for 10 min, rinsed in water, immersed in 60% isopropanol, and then incubated with Oil Red O solution (saturated Oil Red O in isopropanol, diluted 2:3 in 60% isopropanol; Sigma-Aldrich, St. Louis, MO, USA) at room temperature for 40 min. Slides were subsequently washed twice in 60% isopropanol and counterstained with hematoxylin (DAKO, Carpinteria, CA, USA) for 30 s.

2.6. Immunohistochemical Staining

Immunohistochemical staining was performed to evaluate the expression of cluster of differentiation 68 (CD68), lymphocyte antigen 6 complex, locus G (Ly6G), tumor necrosis factor-alpha (TNFα), collagen type I alpha 1 chain (Col1a1), proliferating cell nuclear antigen (PCNA) and antigen Kiel 67 (Ki67). Deparaffinized and rehydrated liver sections were incubated with 0.3% hydrogen peroxide for 30 min to quench endogenous peroxidase activity, followed by washing with distilled water. Antigen retrieval was performed in Tris-ethylenediaminetetraacetic acid (EDTA) buffer at 95 °C for 15 min. After cooling to room temperature, sections were blocked with 5% bovine serum albumin (BSA) for 60 min at room temperature. Sections were subsequently incubated overnight at 4 °C in a humidified chamber with the following primary antibodies diluted in 5% BSA: rabbit anti-mouse CD68 (1:1000, Abcam, Waltham, MA, USA), Ly6G (1:200, Cell Signaling Technology, Danvers, MA, USA), TNFα (1:200, Proteintech, Rosemont, IL, USA), Col1a1 (1:200, Cell Signaling Technology, Danvers, MA), PCNA (1:2000, Cell Signaling Technology, Danvers, MA) or Ki67 (1:4000, Proteintech, Rosemont, IL, USA). After three washes with TBS containing 0.1% Tween 20 (TBST), sections were incubated with horseradish peroxidase-conjugated goat anti-rabbit IgG and visualized using 3,3-diaminobenzidine kit (Vector Laboratories, Inc., Newark, CA, USA) according to the manufacturer’s instructions. Sections were then counterstained with hematoxylin. Quantification was performed by counting CD68+, Ly6G+, PCNA+, and Ki67+ cells per 40× field. Positive staining areas for TNFα and Col1a1 were measured per 40× field using Image J version 1.54p (NIH, Bethesda, MD, USA).

2.7. RNA Extraction, cDNA Synthesis and Quantitative Real-Time Reverse Transcription PCR (RT-qPCR)

Total RNA was extracted from liver tissue using TRIzol reagent (Invitrogen, Carlsbad, CA, USA). RNA concentration was measured using a Nanodrop ND-1000 spectrophotometer, and 1 μg of total RNA was treated to remove genomic DNA and subsequently used for first-strand complementary DNA (cDNA) synthesis with a reverse transcription kit (ABclonal Technology, Woburn, MA, USA) following the manufacturer’s instructions. RT-qPCR was performed on a QuantStudio™ 3 Real-Time PCR System (Applied Biosystems, Thermo Fisher Scientific, Grand Island, NY, USA) using either the SYBR Green PCR Master Mix kit or Probe Master Mix (ABclonal Technology, Woburn, MA, USA) following the manufacturer’s protocols. Gene expression levels were normalized to Rn18s. All TaqMan® assay-on-demand primers were obtained from Thermo Fisher Scientific Inc. (Grand Island, NY, USA) with the following assay IDs: Rn18s, Mm04277571; carnitine palmitoyltransferase 1α (CPT1α), Mm01231183; collagen type I alpha 2 chain (Col1a2), Mm004083888; transforming growth factor-beta (TGFβ), Mm01178820; TNFα, Mm00443258; chemokine (C-C motif) ligand 2 (CCL2), Mm00441242; interleukin 1 beta (IL1b), Mm00434228; intercellular adhesion molecule (ICAM), Mm00516023; connective tissue growth factor (CTGF), Mm01192933. The hFGF1ΔHBS TaqMan primer was custom-designed by Thermo Fisher Scientific based on the previously described sequence [20]. SYBR Green primers were purchased from Sigma-Aldrich (St. Louis, MO, USA) and the sequences were as follows (5′ → 3′): Rn18s, AGTCCCTGCCCTTTGTACACA, CGATCCGAGGGCCTCACTA; mouse FGF1 (mFGF1), GGGGAGATCACAACCTTCGC, GTCCCTTGTCCCATCCACG; CXCL10, CCAAGTGCTGCCGTCATTTTC, GGCTCGCAGGGATGATTTCAA; Col1a1, GCTCCTCTTAGGGGCCACT, CCACGTCTCACCATTGGGG; collagen type XIII alpha 1 chain (Col13a1), CTGTAACATGGAAACTGGGGAAA, CCATAGCTGAACTGAAAACCACC; stearoyl-CoA desaturase-1 (SCD1), GCGAGGGCTTCCACAACTAC, GGCAGCCATGCAGTCGATGA; cluster of differentiation 36 (CD36), ATGGGCTGTGATCGGAACTG, TTTGCCACGTCATCTGGGTTT; Desmin, CCTGGAGCGCAGAATCGAAT, TGAGTCAAGTCTGAAACCTTGGA.

2.8. Western Blot Analysis

Liver tissues were homogenized in Radioimmunoprecipitation Lysis Buffer System (Santa Cruz Biotechnology, Dallas, TX, USA) at 4 °C for 30 min, followed by centrifugation at 13,000 g at 4 °C for 30 min. The lipid layer was removed, and the clear supernatant was collected. Protein concentration was determined using the Protein Assay Dye Reagent Concentrate (Bio-Rad, Hercules, CA, USA). A total of 10–20 µg protein was separated by 10–12% sodium dodecyl sulfate polyacrylamide gel electrophoresis and transferred onto a 0.22 µm nitrocellulose membrane. After transfer, membranes were blocked with 5% non-fat milk or BSA for 1–2 h at room temperature and then incubated overnight at 4 °C with primary antibodies against FGF1 (1:1000, Abcam, Waltham, MA, USA), Zoanthus sp. green fluorescent protein (ZsGreen) (1:1000, OriGene Technologies, Rockville, MD, USA), Flag (1:10,000, Proteintech, Rosemont, IL, USA), SCD1 (1:5000, Proteintech, Rosemont, IL, USA), CD36 (1:1000, Cell Signaling Technology, Danvers, MA, USA), CPT1α (1:5000, Proteintech, Rosemont, IL, USA), β-actin (1:20,000, Proteintech, Rosemont, IL, USA). Membranes were washed three times with TBST and then incubated with peroxidase conjugated goat anti-rabbit or anti-mouse IgG secondary antibodies (1:2000–6000, Cell Signaling Technology, Danvers, MA, USA) at room temperature for 1 h. After three additional washes with TBST, protein bands were visualized using enhanced chemiluminescence reagent (Bio-Rad, Hercules, CA) and imaged with a ChemiDocTM MP Imaging System (Bio-Rad, Hercules, CA, USA). Band densities were quantified using ImageJ version 1.54p (NIH, Bethesda, MD, USA), with β-actin serving as a loading control.

2.9. Statistical Analysis

Data are presented as means ± SEM and were analyzed using GraphPad Prism version 10 software. For comparisons among multiple groups, one-way or two-way ANOVA followed by Tukey’s multiple comparison test was performed. For comparisons between two independent groups, an unpaired two-tailed Student’s t-test was used if the data were normally distributed, whereas the Mann–Whitney U test was applied for non-normally distributed data. Differences were considered statistically significant at p < 0.05.

3. Results

3.1. Induction of MASH in Mice and Experimental Design for Liver-Directed hFGF1ΔHBS Overexpression

To evaluate the therapeutic efficacy of liver-specific overexpression of hFGF1ΔHBS in MASH, 10-week-old ApoE-KO mice were fed a HFHC diet for 4 weeks, followed by tail-vein administration of AAV8-hFGF1ΔHBS (Figure 1A). Mice were maintained on the HFHC diet for an additional 8 weeks after viral delivery. Following IPGTT, IPITT, and DEXA body composition analysis within the last 2 weeks, mice were euthanized for tissue collection. Chow-fed control and HFHC-fed control mice received AAV8-Control vectors.
The efficiency of hepatic hFGF1ΔHBS overexpression was validated by RT-qPCR and Western blot analyses. RT-qPCR analysis showed that hFGF1ΔHBS mRNA was undetectable in AAV8-Control-treated mice, whereas mice receiving AAV8-hFGF1ΔHBS exhibited a clear dose-dependent increase in hepatic hFGF1ΔHBS mRNA expression across different viral doses (Figure 1B). Consistent with the mRNA data, Western blot analysis confirmed the absence of hFGF1ΔHBS protein in AAV8-Control mice, while robust hepatic overexpression of hFGF1ΔHBS protein was observed in mice treated with AAV8-hFGF1ΔHBS in a dose-dependent manner (Figure 1D,E). ZsGreen, used as a viral reporter gene [26], was detectable in all groups (Figure 1D). The expression pattern and band localization of the FLAG tag were consistent with those of hFGF1ΔHBS (Figure 1D), further confirming graded hepatic overexpression of the target gene hFGF1ΔHBS.
RT-qPCR analysis revealed no significant differences in endogenous mFGF1 mRNA levels among groups (Figure 1C). In contrast, mFGF1 protein levels were markedly reduced in AAV8-Control-treated HFHC-fed mice but were restored following hFGF1ΔHBS gene transfer (Figure 1D,F). Notably, mFGF1 protein levels did not differ among the different AAV8-hFGF1ΔHBS dose groups, indicating that hFGF1ΔHBS-mediated restoration of endogenous mFGF1 protein occurs independently of viral dose. This dose-independent restoration, despite unchanged mRNA levels, suggests that hFGF1ΔHBS may normalize endogenous mFGF1 protein expression through post-transcriptional mechanisms.

3.2. Liver-Directed Overexpression of hFGF1ΔHBS Shows No Effect on Systemic Metabolism

Body weight analysis showed that ApoE-KO mice fed a HFHC diet gained significantly more weight than those maintained on a chow diet after 4 weeks. However, no significant differences in body weight were observed between HFHC-fed ApoE-KO mice receiving AAV8-Control or AAV8-hFGF1ΔHBS (Figure 2A). Blood glucose levels were comparable among all groups (Figure 2B). IPGTT and IPITT were performed to evaluate systemic glucose tolerance and insulin sensitivity, respectively, and revealed no significant differences between HFHC-fed ApoE-KO mice with or without AAV8-hFGF1ΔHBS treatment (Figure 2C–F). Consistent with these findings, DEXA analysis showed no differences in total fat mass or lean mass between HFHC-fed ApoE-KO mice regardless of hepatic hFGF1ΔHBS expression (Figure 2G–J). Furthermore, plasma triglyceride and total cholesterol were not affected by liver-directed overexpression of hFGF1ΔHBS in HFHC-fed mice (Figure 2K,L). Collectively, these findings demonstrate that liver-specific overexpression of hFGF1ΔHBS does not alter systemic metabolic parameters under HFHC diet feeding conditions.

3.3. Liver-Directed Overexpression of hFGF1ΔHBS Attenuates Hepatic Lipid Accumulation

Lipid accumulation is a central pathophysiological feature of MASH [27]. Previously, our group demonstrated that recombinant FGF1ΔHBS ameliorates hepatic lipid disorder in db/db and ApoE-KO mice [21]. Building on these findings, we first evaluated the effects of liver-directed hFGF1ΔHBS overexpression on hepatic lipid accumulation.
HFHC diet feeding slightly increased liver size and weight; however, no significant differences were observed between HFHC-fed mice with or without AAV8-hFGF1ΔHBS treatment (Figure 3A–C). In contrast, histological analysis using H&E and Oil Red O staining revealed a marked improvement in hepatic morphology following hFGF1ΔHBS overexpression, characterized by a substantial reduction in lipid droplet deposition (Figure 3D). Consistent with these observations, hepatic triglyceride content was significantly reduced in mice with liver-directed hFGF1ΔHBS overexpression (Figure 3E), whereas hepatic cholesterol levels were not significantly affected (Figure 3F).
In addition, liver-directed hFGF1ΔHBS delivery significantly reduced plasma markers of liver injury, including ALT (Figure 3G) and AST (Figure 3H). Notably, the reduction in hepatic lipid accumulation did not exhibit dose-dependent effects between the 4 × 109 and 2 × 1010 vg/mouse doses of AAV8-hFGF1ΔHBS.
Collectively, these results indicate that liver-specific overexpression of hFGF1ΔHBS effectively alleviates hepatic lipid accumulation in ApoE-KO mice fed an HFHC diet. Moreover, long-term overexpression of hFGF1ΔHBS did not induce liver morphological abnormalities or injury, suggesting that sustained expression of hFGF1ΔHBS in liver is potentially safe.

3.4. Liver-Directed Overexpression of hFGF1ΔHBS Attenuates Hepatic Inflammation

Inflammation is recognized as a key pathological mechanism of MASH [28], and accumulating evidence indicates that recombinant FGF1 effectively suppresses inflammation in metabolic-related diseases [29,30,31]. To determine whether liver-specific overexpression of hFGF1ΔHBS attenuates hepatic inflammation, we evaluated inflammatory cell infiltration and the expression of pro-inflammatory cytokines. CD68 and Ly6G are well-established cell surface markers for macrophages and neutrophils, respectively. Immunohistochemical analysis revealed that macrophage infiltration was markedly increased in AAV8-Control mice fed HFHC diet but was significantly reduced following AAV8-hFGF1ΔHBS transduction (Figure 4A,B). Similarly, Ly6G staining showed a significant increase in neutrophil infiltration AAV8-Control HFHC-fed mice, which was substantially decreased in AAV8-hFGF1ΔHBS-treated mice (Figure 4A,C). Consistent with these findings, the protein expression of the pro-inflammatory cytokine TNFα was markedly elevated in AAV8-Control HFHC-fed mice but significantly decreased after AAV8-hFGF1ΔHBS treatment (Figure 4A,D). TNFα mRNA expression also showed a significant reduction following hFGF1ΔHBS overexpression (Figure 4E). Furthermore, the mRNA expression of IL-1β (Figure 4F), CXCL10 (Figure 4G), CCL2 (Figure 4H), and ICAM (Figure 4I) was upregulated in AAV8-Control HFHC-fed mice but markedly decreased after AAV8-hFGF1ΔHBS transduction. These results demonstrate that liver-directed overexpression of hFGF1ΔHBS effectively suppresses hepatic inflammation. Notably, the anti-inflammatory effect of hFGF1ΔHBS did not show a dose-dependent pattern across the different AAV8-hFGF1ΔHBS doses.

3.5. Liver-Directed Overexpression of hFGF1ΔHBS Attenuates Hepatic Fibrosis

Fibrosis represents the strongest histological predictor of disease progression and clinical outcomes in MASH [32]. To determine whether liver-specific overexpression of hFGF1ΔHBS attenuates hepatic fibrosis, collagen deposition was evaluated by Sirius Red and Masson’s trichrome staining. The results showed that collagen deposition in AAV8-hFGF1ΔHBS-treated HFHC-fed mice was markedly reduced compared with that in AAV8-Control-treated HFHC-fed mice (Figure 5A–C). Col1a1, a well-established marker of hepatic stellate cell activation and liver fibrosis, was further examined by immunohistochemical staining. Compared with AAV8-Control-treated HFHC-fed mice, HFHC-fed mice treated with AAV8-hFGF1ΔHBS exhibited significantly lower Col1a1-positive staining (Figure 5A,D). Consistent with these findings, Col1a1 mRNA expression was significantly reduced in AAV8-hFGF1ΔHBS-treated HFHC-fed groups (Figure 5E). Moreover, the mRNA expression levels of key profibrotic genes, including CTGF (Figure 5F), TGFβ (Figure 5G), Col1a2 (Figure 5H), Col13a1 (Figure 5I), and Desmin (Figure 5J), were all significantly lower in AAV8-hFGF1ΔHBS-treated HFHC-fed mice than in AAV8-Control-treated HFHC-fed mice. Together, these data demonstrate that liver-directed overexpression of hFGF1ΔHBS effectively attenuates hepatic fibrosis. Notably, the antifibrotic effect of hFGF1ΔHBS did not show a clear dose-dependent response across the different AAV8-hFGF1ΔHBS doses tested.

3.6. Liver-Directed Overexpression of hFGF1ΔHBS Inhibits Hepatic Lipid Synthesis and Uptake

As noted above, hepatic delivery of hFGF1ΔHBS reduced lipid accumulation in HFHC-fed mice. In addition, our group previously demonstrated that recombinant FGF1ΔHBS corrected lipid dysregulation and ameliorated fatty liver disease [21]. Based on these observations, we investigated whether hFGF1ΔHBS transfer regulates the expression of genes involved in hepatic fatty acid metabolism in MASH. Liver-directed overexpression of hFGF1ΔHBS reduced hepatic lipid accumulation without a dose-dependent response across different AAV8-hFGF1ΔHBS doses. Therefore, only the high-dose AAV8-hFGF1ΔHBS group was selected to assess the mechanisms underlying lipid accumulation. First, we examined the expression of SCD1, a key enzyme that promotes lipogenesis and triglyceride synthesis. SCD1 mRNA expression was markedly upregulated in AAV8-Control-treated HFHC-fed mice and was significantly reversed following hFGF1ΔHBS transfer (Figure 6A). Consistently, SCD1 protein levels were lower in AAV8-hFGF1ΔHBS-treated HFHC-fed mice compared with AAV8-Control-treated HFHC-fed mice (Figure 6D,E). We next examined the expression of CD36, a fatty acid transporter that mediates long-chain fatty acid uptake. Both CD36 mRNA (Figure 6B) and protein (Figure 6D,F) levels were increased in AAV8-Control-treated HFHC-fed mice, whereas hFGF1ΔHBS transfer markedly reversed these changes. Unexpectedly, mRNA expression of CPT1α, the rate-limiting enzyme of mitochondrial fatty acid β-oxidation, was slightly increased by HFHC diet feeding and reduced following hFGF1ΔHBS transfer, while CPT1α protein levels did not differ among groups (Figure 6C,D,G). Taken together, these findings suggest that liver-directed overexpression of hFGF1ΔHBS ameliorates hepatic lipid accumulation primarily by suppressing lipid uptake and fatty acid synthesis.

3.7. Long-Term Liver-Directed Overexpression of hFGF1ΔHBS Does Not Induce Hepatic Hyperproliferation

Wild-type FGF1 exhibits potent mitogenic activity, raising safety concerns regarding its therapeutic application. To address this limitation, a non-mitogenic FGF1ΔHBS variant has been developed, which retains metabolic benefits while exhibiting markedly reduced mitogenic activity. However, much of this existing evidence supporting the safety of FGF1ΔHBS comes from studies using recombinant protein. Therefore, we evaluated the safety of long-term hepatic overexpression of hFGF1ΔHBS. Hepatocyte proliferation was assessed using PCNA and Ki67 as proliferation markers. Immunohistochemical staining revealed no increase in the numbers of PCNA- and Ki67-positive cells following hFGF1ΔHBS gene transfer (Figure 7A,B). These results indicate that long-term hepatic overexpression of hFGF1ΔHBS does not promote hepatocyte proliferation and supports its safety for therapeutic use.

4. Discussion

The present study provides the first evidence that a single administration of an AAV vector encoding hFGF1ΔHBS confers durable protection against MASH in ApoE-KO mice. We demonstrate that liver-directed hFGF1ΔHBS expression markedly ameliorates key pathological features of MASH, including hepatic lipid accumulation, inflammation, and fibrosis, thereby protecting against disease progression. Importantly, these hepatoprotective effects occurred in the absence of detectable hepatocyte proliferation, supporting the lack of mitogenic risk associated with long-term overexpression of the hFGF1ΔHBS variant. To our knowledge, this represents the first long-term evaluation of liver-restricted hFGF1ΔHBS gene therapy in HFHC-fed mice and demonstrates both the efficacy and safety of this therapeutic strategy.
ApoE-KO mice were selected because ApoE deficiency leads to severe hypercholesterolemia and markedly increases hepatic susceptibility to cholesterol overload. Hepatic accumulation of free cholesterol is a key driver of lipotoxicity, Kupffer cell activation, and inflammasome signaling, processes that collectively promote the development of steatohepatitis [33]. When challenged with a HFHC diet, ApoE-KO mice consistently develop hepatic steatosis, inflammation, hepatocellular ballooning, and early fibrosis, closely recapitulating the histopathological hallmarks of human MASH. This model therefore provided a stringent platform to evaluate the therapeutic efficacy of hepatocyte-directed hFGF1ΔHBS expression under conditions of pronounced hepatic lipotoxic stress that closely mirror human disease progression.
The key novel finding of this study is that the beneficial effects of hFGF1ΔHBS on MASH occur independently of improvements in systemic metabolic parameters. The protective effects of FGF1ΔHBS in metabolic syndrome and its associated complications, including diabetic cardiomyopathy [34], diabetic nephropathy [35,36,37], and fatty liver disease [21], have been extensively investigated using systemically administered recombinant proteins. In these studies, FGF1ΔHBS consistently ameliorated disease pathology; however, these improvements were invariably accompanied by marked correction of systemic metabolic disturbances, leaving unresolved the question of whether FGF1ΔHBS exerts direct organ-specific effects in vivo.
For example, in models of diabetic cardiomyopathy, hFGF1ΔHBS improved mitochondrial integrity and reduced reactive oxygen species accumulation in cardiac tissue while simultaneously alleviating hyperinsulinemia and improving glucose tolerance in db/db mice. Although complementary in vitro evidence in neonatal cardiomyocytes supports direct cardioprotective effects, definitive in vivo evidence for cardiac-specific actions remains limited [34]. Similarly, in diabetic nephropathy models, hFGF1ΔHBS attenuated renal pathology in parallel with substantial reductions in blood glucose levels, while mechanistic investigations were largely confined to podocytes [35,37] or proximal tubule cells [36] in vitro. Consequently, whether renal protection resulted from direct renal action or secondary metabolic improvements remained unclear. Comparable limitations apply to studies of fatty liver disease, in which recombinant FGF1ΔHBS ameliorated steatohepatitis in db/db and ApoE-KO mice while concurrently improving insulin resistance and circulating lipid profiles, with liver-specific mechanisms inferred primarily from hepatocyte culture studies [21].
Notably, a study employing native FGF1 recombinant protein reported amelioration of type 1 diabetic nephropathy without affecting body weight or systemic glycemia [31]. Similarly, Zhang et al. demonstrated that FGF1ΔHBS prevented diabetic cardiomyopathy and hypertension in type 1 diabetic mice, again without altering hyperglycemia or body weight [38]. These findings indicate that FGF1-based therapies can exert protective effects independently of systemic metabolic normalization. However, neither study [31,38] provided evidence for direct liver-specific targeting in vivo. Building on this gap, our study specifically examined the effects of hepatocyte-restricted hFGF1ΔHBS expression in MASH to determine whether FGF1ΔHBS can ameliorate fatty liver disease independently of systemic metabolic improvement. Our results demonstrate that liver-specific hFGF1ΔHBS expression does not alter body weight, glycemia, lipid metabolism, glucose tolerance, or insulin sensitivity in ApoE-KO mice, yet markedly attenuates hepatic lipid accumulation, inflammation, and fibrosis. These findings provide direct in vivo evidence that hFGF1ΔHBS confers liver-intrinsic therapeutic benefits in MASH and suggest that the protective actions of FGF1ΔHBS observed in other metabolic complications may likewise involve tissue-specific mechanisms rather than global metabolic correction.
Although the cellular targets and putative mechanism of action of FGF1ΔHBS in liver remain unclear, multiple lines of evidence point to hepatocytes as the initiating cellular target of hFGF1ΔHBS in vivo. Specifically in this study, the gene-delivery strategy itself strongly biases expression to hepatocytes: AAV8 exhibits pronounced hepatotropism following systemic administration, and the TBG promoter drives hepatocyte-selective transcription. Consistent with this design, we observed robust, dose-responsive increases in hepatic hFGF1ΔHBS mRNA and protein in AAV8–hFGF1ΔHBS-treated mice, supporting hepatocytes as the principal “factory” for transgene synthesis and secretion. Furthermore, our prior in vitro work in primary hepatocytes demonstrated that recombinant hFGF1ΔHBS directly rescues palmitate-induced lipid accumulation [21], aligning with the in vivo reduction in hepatic triglycerides and the downregulation of SCD1 and CD36 observed here. Together, these data suggest that direct actions on hepatocyte lipid handling are sufficient to account for the improvement in steatosis.
At the same time, FGF1 is classically considered a paracrine factor [6], and hepatocyte-derived hFGF1ΔHBS likely engages non-parenchymal cells within the hepatic microenvironment. We therefore posit a model in which secreted hFGF1ΔHBS attenuates Kupffer cell activation and neutrophil recruitment (consistent with reduced CD68 and Ly6G signals, and lower TNFα, CCL2, and CXCL10) while indirectly limiting hepatic stellate cell activation and collagen deposition (reduced Col1a1 protein and transcript, and lower CTGF, TGFβ, Col1a2, Col13a1, and Desmin). In this framework, hepatocytes are the primary initiators through correction of lipid metabolic stress, and paracrine crosstalk secondarily dampens inflammatory and fibrogenic programs, together producing the composite histologic benefit observed in MASH.
A major safety concern associated with chronic hFGF1ΔHBS therapy is the potential risk of tumorigenesis, given the potent mitogenic activity of native FGF1 [39]. FGF1ΔHBS was engineered to preserve the metabolic actions of FGF1 while markedly reducing its proliferative activity, a concept supported by multiple in vivo studies [20,21]. We previously demonstrated that repeated intraperitoneal (0.5 mg/kg every other day for up to 3 months) or intravenous (2.0 mg/kg every other day for up to 1 month) administration of FGF1ΔHBS did not induce hepatic proliferation in healthy C57BL/6J mice [20]. However, metabolic liver diseases such as fatty liver and MASH represent preneoplastic conditions in which proliferative safety requires more rigorous evaluation.
Consistent with prior studies in healthy mice, long-term systemic administration of recombinant FGF1ΔHBS (0.5 mg/kg every other day for up to 3 months) did not increase hepatic [21] and renal [35,37] cell proliferation in db/db mice, supporting its favorable safety profile under protein-based treatment paradigms. However, these studies involved intermittent systemic delivery of a short-lived recombinant protein and therefore do not necessarily predict the consequences of sustained, liver-directed transgene expression. In the present study, long-term hepatic expression of hFGF1ΔHBS did not alter liver size or weight and did not increase PCNA or Ki67 expression in HFHC-fed ApoE-KO mice, confirming the safety profile of hFGF1ΔHBS even under conditions of persistent hepatocyte-specific overexpression.
A recent work [40] demonstrated that under conditions of extreme systemic exposure, daily administration of a very high dose of recombinant hFGF1ΔHBS (3 mg/kg) for 12 consecutive days, residual proliferative signaling could still be detected in proliferation-sensitive tissues such as the bladder. Strikingly, fusion of FGF1ΔHBS to a muscle-targeting peptide eliminated aberrant proliferative signaling while preserving metabolic efficacy, underscoring the critical importance of tissue-restricted delivery as a decisive determinant of safety [40]. These findings provide strong mechanistic support for our liver-directed AAV strategy, which confines hFGF1ΔHBS expression to hepatocytes, minimizes systemic exposure, and avoids unintended FGF receptor activation in extrahepatic tissues.
Beyond the safety engineering of the ligand itself, the AAV8 vector provides a second layer of safety. First, the AAV8 serotype exhibits strong hepatotropism, which naturally confines transgene expression to the liver and minimizes off-target expression in extrahepatic tissues [24,41]. Second, recombinant AAV genomes exist predominantly as non-replicating episomes within the nucleus rather than integrating into the host genome, thereby minimizing the risk of insertional mutagenesis [42]. The clinical safety of AAV8-mediated liver-directed gene transfer has been well established in human trials for hemophilia B [43,44]. Together, our data suggest that the combination of the hFGF1ΔHBS and the non-integrating, liver-specific AAV8 vector offers a potentially safe and effective approach for MASH treatment.
Currently approved pharmacologic therapies for MASH with metabolic syndrome, including agents such as Resmetirom, a thyroid hormone receptor-β agonist [45], and GLP-1 receptor agonists such as semaglutide [46], primarily target systemic metabolic dysregulation to achieve secondary hepatic benefits. These agents are effective in obese or diabetic patients but require continuous, often lifelong administration and are associated with dose-limiting gastrointestinal or cardiovascular side effects [45,46]. In contrast, AAV8-hFGF1ΔHBS gene therapy offers a fundamentally different, liver-targeted strategy that acts directly on hepatocyte lipid metabolism and inflammatory pathways without altering body weight, glucose homeostasis, or circulating lipids. This metabolic independence suggests particular relevance for lean MASH, a growing patient subgroup in whom systemic metabolic therapies may be unnecessary or less effective [47]. By enabling sustained, hepatocyte-restricted expression after a single administration, AAV8-hFGF1ΔHBS may therefore provide a precision therapeutic modality that circumvents the limitations of chronic systemic therapy while directly addressing the liver-intrinsic mechanisms driving disease progression in lean MASH.
Beyond metabolic liver disease, liver-restricted hFGF1ΔHBS expression may also hold therapeutic potential in other forms of liver injury, including anti-tuberculosis drug-induced liver injury. A previous study showed that isoniazid- and rifampicin-induced liver injury is associated with reduced hepatic FGF1 expression, and that recombinant hFGF1ΔHBS ameliorates liver injury [48]. However, systemic delivery of hFGF1ΔHBS may be undesirable in patients with active tuberculosis, where excessive metabolic modulation or unintended weight loss could be detrimental. Liver-specific expression of hFGF1ΔHBS therefore represents a rational alternative that preserves hepatoprotective efficacy while minimizing systemic effects.
Several limitations should be acknowledged. First, the ApoE-KO HFHC model reflects a cholesterol-driven form of steatohepatitis and does not encompass the full heterogeneity of human MASH. Validation of therapeutic efficacy in additional models, including diet-induced obesity, hypertension-associated MASH, and diabetes-associated fatty liver diseases, will be essential to generalize our findings. Second, the current study primarily evaluates early intervention rather than treatment of advanced disease. Although our prior work demonstrated therapeutic efficacy of recombinant hFGF1ΔHBS in severely diabetic mice [21], dedicated studies using liver-directed hFGF1ΔHBS in advanced MASH models are still required. Third, evaluation in lean MASH models will be critical to define the translational relevance of this approach across patient subtypes. Finally, AAV8-mediated gene delivery results in durable but largely irreversible hepatic expression; accordingly, comprehensive long-term safety and efficacy studies in large-animal models, including non-human primates, are required to support the clinical translational potential of AAV-hFGF1ΔHBS-based gene therapy for these highly prevalent diseases.

5. Conclusions

Liver-directed overexpression of hFGF1ΔHBS ameliorates MASH in ApoE-KO mice independently of systemic metabolic regulation, significantly reducing hepatic inflammation, fibrosis, and lipid accumulation through coordinated suppression of lipid synthesis and uptake. Importantly, these therapeutic effects are achieved without evidence of increased hepatocyte proliferation, supporting the safety of the sustained expression of this non-mitogenic FGF1 variant. Collectively, our findings establish liver-specific hFGF1ΔHBS gene therapy as a promising and mechanistically targeted strategy for the treatment of MASH.

Author Contributions

Conceptualization, Y.T. and Y.L.; methodology, Y.L., X.H., C.G., Q.L., A.A.-I., Z.X., S.Y., H.H. and J.X.; validation, Y.T. and Y.L.; formal analysis, Y.T., Y.L. and X.H.; investigation, Y.L. and X.H.; resources, Y.T., K.A.W., L.C. and Z.D.; data curation, Y.L., X.H. and Y.T.; writing—original draft preparation, Y.T., and Y.L.; writing—review and editing, Y.T., K.A.W., S.E.W., L.C. and Z.D.; visualization, Y.T. and Y.L.; supervision, Y.T. and Z.D.; project administration, Y.T. and Z.D.; funding acquisition, Y.T., L.C. and Z.D. All authors have read and agreed to the published version of the manuscript.

Funding

This study was supported in part by R01 grants from the National Institutes of Health (R01HL125877, R01HL160927, and R01HL174922), Kentucky Pediatric Cancer Research Trust Fund FY 24/25, and the Jewish Heritage Fund for Excellence Research Enhancement Grant Program at the University of Louisville School of Medicine to Y.T. D.Z. is supported in part by NIH R01DK131442 and R01DK115406, and L.C. is supported in part by NIH P30ES030328.

Institutional Review Board Statement

All animal procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Louisville under protocol numbers 18189 and 24400, approved on 26 August 2022 and 5 August 2024, respectively.

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in this study are included in the article. Further inquiries can be directed to the corresponding authors.

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 1. Induction of a MASH mouse model and experimental design for AAV8-hFGF1ΔHBS intervention. ApoE-KO mice were fed a HFHC diet for 4 weeks and then administered AAV8-hFGF1ΔHBS at doses of 4 × 109 or 2 × 1010 vg/mouse. Chow-fed control mice received 2 × 1010 vg of AAV8-Control. The mice received additional 10 weeks HFHC diet or chow. (A) Schematic illustration of the experimental timeline, including HFHC diet feeding, virus injection, IPGTT, IPITT and DEXA scan. After euthanasia, liver tissues were collected and (B,C) relative mRNA of hFGF1ΔHBS and mFGF1 were detected by RT-qPCR, with 18s rRNA used as housekeeping gene; (D–F) protein expression of hFGF1ΔHBS, mFGF1, ZsGreen, and Flag were detected by Western blot, with β-actin used as the loading control, and quantified by densitometry. FGF1-KO mice were used as a negative control to identify mFGF1 band, and different exposure (Ex.) times were applied to distinguish hFGF1ΔHBS and mFGF1 bands. Data are presented as means ± SEM (n = 3–7). Statistical significance was defined as p < 0.05.
Figure 1. Induction of a MASH mouse model and experimental design for AAV8-hFGF1ΔHBS intervention. ApoE-KO mice were fed a HFHC diet for 4 weeks and then administered AAV8-hFGF1ΔHBS at doses of 4 × 109 or 2 × 1010 vg/mouse. Chow-fed control mice received 2 × 1010 vg of AAV8-Control. The mice received additional 10 weeks HFHC diet or chow. (A) Schematic illustration of the experimental timeline, including HFHC diet feeding, virus injection, IPGTT, IPITT and DEXA scan. After euthanasia, liver tissues were collected and (B,C) relative mRNA of hFGF1ΔHBS and mFGF1 were detected by RT-qPCR, with 18s rRNA used as housekeeping gene; (D–F) protein expression of hFGF1ΔHBS, mFGF1, ZsGreen, and Flag were detected by Western blot, with β-actin used as the loading control, and quantified by densitometry. FGF1-KO mice were used as a negative control to identify mFGF1 band, and different exposure (Ex.) times were applied to distinguish hFGF1ΔHBS and mFGF1 bands. Data are presented as means ± SEM (n = 3–7). Statistical significance was defined as p < 0.05.
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Figure 2. AAV8-hFGF1ΔHBS treatment does not significantly alter systemic metabolic parameters. ApoE-KO mice were fed and treated as described in Figure 1. (A) Body weight and (B) blood glucose levels were monitored over time. During the final two weeks of the study, (C) intraperitoneal glucose tolerance test (IPGTT) and (D) area under the curve (AUC) of blood glucose during IPGTT, as well as (E) intraperitoneal insulin tolerance test (IPITT) and (F) area under the curve (AUC) of blood glucose during IPITT, were assessed. Body composition was analyzed by dual-energy X-ray absorptiometry (DEXA) scan, including (G) lean mass (%), (H) lean mass (g), (I) fat mass (%), and (J) fat mass (g). After euthanasia, plasma was collected for measurement of (K) triglycerides and (L) total cholesterol. Data are presented as means ± SEM (n = 6–7). Statistical significance was defined as p < 0.05. * p < 0.05 for Chow AAV8-Control vs. HFHC AAV8-Control.
Figure 2. AAV8-hFGF1ΔHBS treatment does not significantly alter systemic metabolic parameters. ApoE-KO mice were fed and treated as described in Figure 1. (A) Body weight and (B) blood glucose levels were monitored over time. During the final two weeks of the study, (C) intraperitoneal glucose tolerance test (IPGTT) and (D) area under the curve (AUC) of blood glucose during IPGTT, as well as (E) intraperitoneal insulin tolerance test (IPITT) and (F) area under the curve (AUC) of blood glucose during IPITT, were assessed. Body composition was analyzed by dual-energy X-ray absorptiometry (DEXA) scan, including (G) lean mass (%), (H) lean mass (g), (I) fat mass (%), and (J) fat mass (g). After euthanasia, plasma was collected for measurement of (K) triglycerides and (L) total cholesterol. Data are presented as means ± SEM (n = 6–7). Statistical significance was defined as p < 0.05. * p < 0.05 for Chow AAV8-Control vs. HFHC AAV8-Control.
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Figure 3. AAV8-hFGF1ΔHBS treatment attenuates lipid accumulation. ApoE-KO mice were fed and treated as described in Figure 1. Following euthanasia, (A) liver size, (B) liver weight, and (C) the percentage of liver weight to body weight were measured. Liver sections were subjected to (D) hematoxylin–eosin (H&E, upper panel) staining and Oil Red O (red color, middle and lower panel) staining. Hepatic contents of (E) triglycerides and (F) total cholesterol were quantified. Plasma markers of liver injury, (G) ALT and (H) AST, were measured. Data are presented as means ± SEM (n = 6–7). Statistical significance was defined as p < 0.05.
Figure 3. AAV8-hFGF1ΔHBS treatment attenuates lipid accumulation. ApoE-KO mice were fed and treated as described in Figure 1. Following euthanasia, (A) liver size, (B) liver weight, and (C) the percentage of liver weight to body weight were measured. Liver sections were subjected to (D) hematoxylin–eosin (H&E, upper panel) staining and Oil Red O (red color, middle and lower panel) staining. Hepatic contents of (E) triglycerides and (F) total cholesterol were quantified. Plasma markers of liver injury, (G) ALT and (H) AST, were measured. Data are presented as means ± SEM (n = 6–7). Statistical significance was defined as p < 0.05.
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Figure 4. AAV8-hFGF1ΔHBS treatment attenuates hepatic inflammation. ApoE-KO mice were fed and treated as described in Figure 1. Liver sections were analyzed by (A–D) immunohistochemical staining to assess macrophage (CD68+, arrowhead) and neutrophil (Ly6G+, arrowhead) infiltration and TNFα expression (arrowhead), along with corresponding quantification. (E–I) Relative mRNA levels of TNFα, IL-1β, CXCL10, CCL2 and ICAM were measured by RT-qPCR, with 18s rRNA used as the housekeeping gene. Data are presented as means ± SEM (n = 6–7). Statistical significance was defined as p < 0.05.
Figure 4. AAV8-hFGF1ΔHBS treatment attenuates hepatic inflammation. ApoE-KO mice were fed and treated as described in Figure 1. Liver sections were analyzed by (A–D) immunohistochemical staining to assess macrophage (CD68+, arrowhead) and neutrophil (Ly6G+, arrowhead) infiltration and TNFα expression (arrowhead), along with corresponding quantification. (E–I) Relative mRNA levels of TNFα, IL-1β, CXCL10, CCL2 and ICAM were measured by RT-qPCR, with 18s rRNA used as the housekeeping gene. Data are presented as means ± SEM (n = 6–7). Statistical significance was defined as p < 0.05.
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Figure 5. AAV8-hFGF1ΔHBS treatment attenuates fibrosis. ApoE-KO mice were fed and treated as described in Figure 1. Liver sections were analyzed by (A–D) Sirius Red (red color, upper panel) and Masson’s trichrome (blue color, middle panel) staining to evaluate collagen deposition, and immunohistochemical staining to assess Col1a1 protein expression (brown color, lower panel), along with corresponding quantification. (E–J) Relative mRNA levels of TGFβ, CTGF, Col1a1, Col1a2, Col13a1 and Desmin were detected by RT-qPCR, 18s rRNA used as the housekeeping gene. Data are presented as means ± SEM (n = 6–7). Statistical significance was defined as p < 0.05.
Figure 5. AAV8-hFGF1ΔHBS treatment attenuates fibrosis. ApoE-KO mice were fed and treated as described in Figure 1. Liver sections were analyzed by (A–D) Sirius Red (red color, upper panel) and Masson’s trichrome (blue color, middle panel) staining to evaluate collagen deposition, and immunohistochemical staining to assess Col1a1 protein expression (brown color, lower panel), along with corresponding quantification. (E–J) Relative mRNA levels of TGFβ, CTGF, Col1a1, Col1a2, Col13a1 and Desmin were detected by RT-qPCR, 18s rRNA used as the housekeeping gene. Data are presented as means ± SEM (n = 6–7). Statistical significance was defined as p < 0.05.
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Figure 6. AAV8-hFGF1ΔHBS treatment improves dysregulated lipid synthesis and transport. ApoE-KO mice were fed and treated as described in Figure 1. (A–C) Relative mRNA levels of SCD1, CPT1α and CD36 were detected by RT-qPCR, 18s rRNA used as the housekeeping gene. (D–G) Protein expression levels of SCD1, CPT1α and CD36 were assessed by Western blot, with β-actin as the loading control, and quantified by densitometric analysis. Data are presented as means ± SEM (n = 5). Statistical significance was defined as p < 0.05.
Figure 6. AAV8-hFGF1ΔHBS treatment improves dysregulated lipid synthesis and transport. ApoE-KO mice were fed and treated as described in Figure 1. (A–C) Relative mRNA levels of SCD1, CPT1α and CD36 were detected by RT-qPCR, 18s rRNA used as the housekeeping gene. (D–G) Protein expression levels of SCD1, CPT1α and CD36 were assessed by Western blot, with β-actin as the loading control, and quantified by densitometric analysis. Data are presented as means ± SEM (n = 5). Statistical significance was defined as p < 0.05.
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Figure 7. AAV8-hFGF1ΔHBS treatment exerts no significant effects on hepatic proliferation. ApoE-KO mice were fed and treated as described in Figure 1. Liver sections were analyzed by (A) immunohistochemical staining for PCNA and Ki67 (arrowhead), along with (B,C) corresponding quantification. Data are presented as means ± SEM (n = 6–7). Statistical significance was defined as p < 0.05.
Figure 7. AAV8-hFGF1ΔHBS treatment exerts no significant effects on hepatic proliferation. ApoE-KO mice were fed and treated as described in Figure 1. Liver sections were analyzed by (A) immunohistochemical staining for PCNA and Ki67 (arrowhead), along with (B,C) corresponding quantification. Data are presented as means ± SEM (n = 6–7). Statistical significance was defined as p < 0.05.
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MDPI and ACS Style

Li, Y.; Hui, X.; Gu, C.; Lin, Q.; Abdelbaset-Ismail, A.; Xu, Z.; Yadav, S.; Huang, H.; Xu, J.; Watson, S.E.; et al. Human FGF1ΔHBS Gene Therapy as Treatment for Metabolic Dysfunction-Associated Steatohepatitis in ApoE-KO Mice. Cells 2026, 15, 387. https://doi.org/10.3390/cells15050387

AMA Style

Li Y, Hui X, Gu C, Lin Q, Abdelbaset-Ismail A, Xu Z, Yadav S, Huang H, Xu J, Watson SE, et al. Human FGF1ΔHBS Gene Therapy as Treatment for Metabolic Dysfunction-Associated Steatohepatitis in ApoE-KO Mice. Cells. 2026; 15(5):387. https://doi.org/10.3390/cells15050387

Chicago/Turabian Style

Li, Yingjian, Xiaodan Hui, Chunjie Gu, Qian Lin, Ahmed Abdelbaset-Ismail, Zixuan Xu, Suchen Yadav, Hongbiao Huang, Jason Xu, Sara E. Watson, and et al. 2026. "Human FGF1ΔHBS Gene Therapy as Treatment for Metabolic Dysfunction-Associated Steatohepatitis in ApoE-KO Mice" Cells 15, no. 5: 387. https://doi.org/10.3390/cells15050387

APA Style

Li, Y., Hui, X., Gu, C., Lin, Q., Abdelbaset-Ismail, A., Xu, Z., Yadav, S., Huang, H., Xu, J., Watson, S. E., Wintergerst, K. A., Cai, L., Deng, Z., & Tan, Y. (2026). Human FGF1ΔHBS Gene Therapy as Treatment for Metabolic Dysfunction-Associated Steatohepatitis in ApoE-KO Mice. Cells, 15(5), 387. https://doi.org/10.3390/cells15050387

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