Introduction

Liver fibrosis is a pathological process with a high prevalence worldwide. It is a characteristic pathological feature of various chronic liver disease and leads to cirrhosis1,2,3. The mechanism of liver fibrosis formation is related to the abnormal activation of hepatic stellate cells (HSCs) and the excessive deposition of extracellular matrix (ECM)4,5. Despite advances in understanding the causes and development of liver fibrosis are clear, there are almost no effective treatments. Early detection and early treatment can intervene in liver fibrosis, but it is still impossible to completely reverse this pathological process6,7. Taking this into account, scholars have been searching for drugs that develop antifibrotic therapies, and have also delved into the molecular mechanisms by which these drugs exert their effects8.

Cuproptosis, as a type of programmed cell death triggered by lipid peroxidation, plays a crucial regulatory role in various fibrotic diseases9,10. The accumulation of copper within cells triggers the aggregation of mitochondrial fatty acylated proteins and the instability of iron-sulfur cluster proteins, leading to cell death and activating inflammatory immune responses11,12. A clinical data report has revealed that copper metabolism disorder is significantly positively correlated with the progression of fibrosis13. Thus, inhibiting the related biological processes of cuproptosis comes to be one of the strategies for treating liver fibrosis. Ferritin 1 (FDX1), along with HSP70 and DLAT, acts as a key regulator of copper ions and are typical markers of cuproptosis. Mechanistically, the downregulation of FDX1 enhances the accumulation of mitochondrial ROS and accelerate cuproptosis14. On the other hand, the activation of the Keap1-Nrf2-Atp7b signaling axis has been proven to be related to the alleviation of cuproptosis15. Merestinib, a specific inhibitor of cuproptosis, was proved to decrease oxidative stress levels by binding directly to NRF2. MTB blocks elesclomol-CuCl2 (ES-Cu) induced cuproptosis by preventing both the aggregation of lipoylated proteins and the destabilization of Fe-S cluster proteins. Nevertheless, cuproptosis as a new research direction for liver fibrosis requires further exploration and investigation16.

Traditional Chinese Medicine (TCM) has demonstrated a therapeutic potential in liver disease management, particularly in addressing inflammatory responses. Studies on the treatment of liver fibrosis with quercetin have confirmed that copper-mediated apoptosis is one of the mechanisms of its pharmacological action17. Recent studies have also shown that Vitisin A, as a type of Pyranoanthocyanin, shows potential value in the treatment of liver diseases. Vitisin A inhibits the ubiquitination of TRAF6 and the formation of the TRAF6-TAK1 complex, blocks the activation of the NF-κB signaling pathway, and reduces the secretion of pro-inflammatory factors such as TNF-α and IL-618. Additionally, Vitisin A regulates liver fat production and fatty acid β-oxidation, which are essential processes for cuproptosis. Furthermore, Vitisin A inhibits 3-hydroxy-3-methylglutaryl coenzyme A reductase (HMGCR) and cholesterol biosynthesis through sterol regulatory element-binding protein 2 (SREBP 2) to inhibiting cholesterol biosynthesis19. This anti-inflammatory and metabolic regulatory effects of Vitisin A may be to regulate copper metabolism by reducing copper ion influx. Vitisin A may down-regulate the expression of the KEAP1/NRF2 pathway, although this requires experimental verification. Moreover, the mechanism of action of Vitisin A also needs to be demonstrated through in vivo and in vitro experiments.

Our current experiments show that Vitisin A inhibits liver fibrosis and cuproptosis in activated LX2s in a time-dependent and dose-dependent manner. Nrf2/HO-1 signaling is the key molecular pathway through which Vitisin A inhibiting liver fibrosis and cuproptosis. In vivo experiments have also demonstrated that Vitisin A has therapeutic effect on fibrosis and cuproptosis by Nrf2/HO-1 signaling pathway. Overall, the aim of this study is to evaluate the antifibrotic effects of Vitisin A and to clarify its molecular mechanism.

Materials and methods

Cell culture and treatment

The immortalized human hepatic stellate cell line LX-2 and human normal hepatocytes MIHA as well as mouse hepatocytes AML12 was purchased from the Cell Bank of Type Culture Collection of Chinese Academy of Sciences (Shanghai, China) and was detected to be free of mycoplasma. LX-2, MIHA, and AML12 cell lines were cultured in Gibco (New York) media: LX-2 in DMEM, MIHA in RPMI 1640, and AML12 in DMEM/F-12. All the three cell medium were supplemented with 10% fetal bovine serum (FBS, Gibco, New York), 100 U/mL penicillin and 100 U/mL streptomycin, and maintained at 37◦C in a humidified environment containing 5% CO2. The culture medium was changed every 48 h. Treatment with TGF-β1 (10 ng/ml in PBS) was used as a positive control and with equivalent PBS as a negative control. Experimental methods, including cell culture and related techniques, followed protocols validated in the co-author’s laboratory and article20. Vitisin A was obtained from ALB Materials (ALB-RS-10075) with a purity of 97.5%. Vitisin A at different dose were added to culture medium over different time to assume the effects on cell lines. ES-Cu (50 nM) was obtained from MedChemExpress (HY-12040) and used as a pharmacological modulator 2 h before treatment where indicated. The si-Nrf2 (5′-CCGAAUUACAGUGUCUUAATT-UUAAGACACUGUAA’UUCGGTT-3′) and siRNA negative control (5′-UUCUCCGAAGUCACGUTT-ACGUGACACGUUCGGAGAATT-3′) were purchased from GenePharma (SuZhou, China) and were transfected into cells using lipofectamine 2000 (Thermo Fisher) for 24 h. The RNA and proteins from cells were collected for quantitative real-time PCR (qRT-PCR), ELISA and western blot analysis.

Animal model induction and treatment

All animal experimental protocols were approved by the institutional review board of Department of Laboratory Animal Science, Anhui Medical University, and conformed in accordance to the National Guidelines for Animal Usage in Research. All animals involving in our study were performed in accordance with ARRIVE guidelines (PLoS Bio 8(6), e1000412,2010)21. All experimental mice were 5–7-week-old males (GemPharmatech LLC, Nanjing, China) housed in a specific-pathogen-free (SPF) animal facility with a 12-hour light/dark cycle, 45–65% humidity, and 20–24 °C temperature. Mice were provided with standard feed, ad libitum water, and bedding changed three times weekly.

In each animal experiment, 6 mice were randomly allocated to each group. To evaluate the roles of Vitisin A in fibrosis model, mice were intraperitoneally injected with 0.5 mL/kg of CCl4 twice a week for a total of 6 weeks. While the control group of mice were given same volume normal saline. Subsequently, Vitisin A in different dose and time were intraperitoneally injected in mice. ML385 (30 mg/kg) was injected peritoneally before Vitisin A treatment. Mice were euthanized via inhalation of isoflurane followed by cervical dislocation, consistent with ARRIVE guidelines. Liver and plasma from mice were collected for next experiment.

CCK8

The viability of cell in different groups was detected using CCK8 assay. Cells were grown in 96-well plates in 100 µl cell culture medium at a density of 4 × 103 cells/well and incubated at 37 °C for 24 h. 10% CCK8 solution was added directly into the culture medium after treatment for the same time. medium after the same time of treatment. Data were subsequently acquired in an enzyme labeller (Bio-Rad, CA) at 570 nm. Three assays were performed in three independent studies and the percentage of activity was determined.

ELISA

Cells were seeded at a density of 4 × 10³ cells per well in a 96-well plate containing 100 µL of cell culture medium and cultured at 37 °C for 24 h to ensure cell adhesion and growth. After treatment, the medium was aspirated, and fresh medium containing specific ELISA detection reagents for all proteins was added. Protein levels of fatty acid synthase (FAS, Human: Abcam, AB279412, Mouse: Reddot Biotech, RD-FASN-Mu), acetyl-CoA carboxylase (ACC, Human: FineTest, EH0481, Mouse: Antibodies, A79069), carnitine palmitoyltransferase 1 (CPT1, Human: MyBioSource, MBS724213, Mouse: FineTest, ER0287), and peroxisome proliferator-activated receptor alpha (PPARα, Human: Antibodies, A74948, Mouse: Biotang, 50–155-351) were quantified using commercially available ELISA kits. We also evaluated expression of copper transporter 1 (CTR1, Human: MyBioSource, MBS9339335, Mouse: Cusabio, CSB-EL015734MO), ATPase copper transporting beta (ATP7B, Human: FineTest, EH15242, Mouse: Elabscience, E-EL-M2918c).

Detection of ROS and MDA levels

Intracellular reactive oxygen species (ROS) levels were assessed using the fluorescent probe 2′,7′-dichlorofluorescein diacetate (DCFH-DA; Beyotime, China). Briefly, cells were seeded in 96-well plates at a density of 4 × 10³ cells/well and cultured at 37 °C for 24 h. Following treatment, cells were incubated with 10 µM DCFH-DA in serum-free medium for 30 min at 37 °C in the dark. Subsequently, cells were washed three times with PBS to remove excess probe, and fluorescence intensity was immediately recorded using a microplate reader (excitation: 488 nm, emission: 525 nm).

Malondialdehyde (MDA), a lipid peroxidation marker, was quantified in both cultured cells and tissue homogenates using commercially available assay kits (Nanjing Jiancheng Bioengineering Institute, China), according to the manufacturer’s protocol. Samples were homogenized and centrifuged to obtain supernatants, which were then incubated with thiobarbituric acid (TBA) reagent under heating conditions. The absorbance of the reaction product was measured spectrophotometrically at 532 nm, and MDA concentration was calculated based on a standard curve and normalized to protein content.

Western blot

Tissues and cells were lysed with strong RIPA buffer (Beyotime, Shanghai) and 1% Phenylmethylsulfonyl fluoride (PMSF, Beyotime, Shanghai) and centrifuged at 4◦C, 12,000 × g for 30 min. The supernatants were collected and the protein concentration was determined by BCA protein assay kit. Total 20 mg proteins in the samples were separated by 10% sodium dodecyl sulfate polyacrylamide gel electrophoresis and transferred to 0.22 μm polyvinylidene difluoride membranes (PVDF, Millipore Corp, Billerica, MA, United States). After blocking in 5% skim milk, the membranes were washed with Tween 20-buffered saline (TBS) and incubated overnight with the specific primary antibodies at 4◦C. Rabbit polyclonal antibodies against α-SMA (1:1000, 14395-1-AP, Proteintech, China), Collagen I (1:1000, ab270993, Abcam, United States), FDX1(1:1000, 12592-1-AP, Proteintech, China), DLAT (1:1000, 13426-1-AP, Proteintech, China), Keap1(1:1000, 10503-2-AP, Proteintech, China), HO-1(1:1000, 10701-1-AP, Proteintech, China), NRF2 (1:1000, 16396-1-AP, Proteintech, China), β-actin (1:5000, 20536-1-AP, Proteintech, China) were used, respectively. The membranes were incubated with the corresponding horseradish peroxidase- conjugated secondary antibodies (anti-mouse or anti-rabbit, Beyotime, Shanghai) for 1 h at room temperature. After washing with TBS, protein bands were visualized with an ECL chemiluminescence kit (ECL-plus, Thermo Fisher, MA, United States) by a chemiluminescent western blotting detection system (chemidoc, BioRad, CA, United States). The intensity of the bands was analyzed with Image J software.

Quantitative real-time PCR

Total RNA was isolated in cell and tissue samples using Trizol reagent (Invitrogen, CA, United States) according to the manufacturer’s instructions. Extracted total RNA was resuspended with RNase-free water and the concentration was measured and using Nano Drop 2,000 (Thermo Fisher, MA, United States). In total RNAs, 500 ng was reverse transcribed to cDNAs and then quantitively analyzed using RT reagent Kit and SYBR-Green Master Mix (Takara, Japan). Primer sequences of Nrf2 is 5’-GGTATTTGACTTCAGTCAA-3’ and 5’-GGCTGAGACTAGTACAGTT-3’. Duplicate of each sample were determined. GAPDH was used as an internal reference. The relative expression of Nrf2 was calculated using the ΔΔCT method.

Hematoxylin-eosin staining and masson staining

Liver tissues from mice were formalin-fixed (10% neutral buffered formalin, 24 h) and paraffin-embedded following routine histological processing; 5-µm sections were stained with H&E (for evaluating inflammatory infiltrates, hepatocellular ballooning, and steatosis) and Masson trichrome (for quantifying collagen deposition as blue-stained fibrosis) using validated protocols. A liver fibrosis specimen was identificated when a large number of blue areas appeared in the section.

Statistical analysis

All data were presented as mean ± standard deviation (SD). Comparisons between two groups were performed using Student’s t-test. For comparisons involving four or more groups, one-way or two-way analysis of variance (ANOVA) combined with Tukey’s post hoc test was used to assess intergroup differences. A p-value < 0.05 was considered statistically significant. All statistical analyses were performed using GraphPad Prism 10.0 software.

Fig. 1
figure 1

Vitisin A inhibits liver fibrosis in a dose-dependtly manner. (A) Representative image of HE staining in mice liver tissue slide. (B) Representative image of Masson staining. (C-F) Expression of four liver fibrosis related serum markers. (G-I) Western blot and quantitative analysis of Collagen I and α-SMA. (J-M) ELISA analysis of cuproptosis related markers. (N) ELISA analysis of critical protein of liver fat production. (O) ELISA analysis of critical markers of fatty acid β-oxidation. (P) MDA relative levels via ELISA analysis. Data are expressed as mean ± SD (n ≥ 5 mice per group). * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001.

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Fig. 2
figure 2

Vitisin A inhibits liver fibrosis and cuproptosis in activated LX2s. (A) Dose-dependent effects of Vitisin A (0–10 nM) on cell viability in normal and fibrotic cells. (B) Time-dependent effects of Vitisin A (0, 2, 4, 6, 8, 12, 24 h) on cell viability in normal and fibrotic cells. (C) Fibrotic cells treated with the cuproptosis activator ES-Cu (0–100 nM), comparing cell viability between ES-Cu-treated fibrotic cells and ES-Cu + Vitisin A co-treated cells. (D-G) ELISA analysis of cuproptosis-related markers including (D) FDX1, (E) DLAT, (F) CTR1, (G) ATP7B. (H-I) Protein expression of fibrosis proteins. (J-K) Detection of critical proteins in (J) liver fat production and (K) fatty acid β-oxidation. (L) ROS levels. (M) MDA levels. Data are expressed as mean ± SD (n ≥ 3 independent experiments). * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001.

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Fig. 3
figure 3

Vitisin A inhibits fibrosis and cuproptosis by Nrf2/HO-1 signaling pathway. (A-C) WB analysis of Nrf2 and HO-1 in Vitisin A and TGF-β1 cultured LX2s. (D) The efficiency of Nrf2 inhibition by siRNA. (E-K) WB analysis of related markers. (L) Quantitative analysis of cuproptosis-related markers. (M) ELISA analysis of liver fat production-related proteins. (N) ELISA analysis of fatty acid β-oxidation-related proteins. (O) ROS levels. (P) MDA levels. Data are expressed as mean ± SD (n ≥ 3 independent experiments). * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001.

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Fig. 4
figure 4

Vitisin A have therapeutic effect on fibrosis and cuproptosis by Nrf2/HO-1 signaling pathway in vivo. (A) Western blot of Nrf2/HO-1 signaling, cuproptosis and fibrosis related proteins in mice treated with Vitisin A. (B) Quantitative analysis of related markers. (C) MDA levels. (D) Liver fat production-related proteins expression. (E) ELISA analysis of fatty acid β-oxidation-related proteins. (F) The efficiency of Nrf2 inhibition by ML385. (G) Quantitative analysis of cuproptosis-related markers. (H) Serum ALT and AST level of mice. (I) Representative image of HE and Masson staining in mice liver tissue. Data are expressed as mean ± SD (n ≥ 5 mice per group). * p < 0.05, ** p < 0.001, *** p < 0.001, **** p < 0.0001.

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Results

Vitisin A inhibits liver fibrosis in a dose-dependtly manner

To investigate the effect of Vitisin A on liver fibrosis in vivo, chronic liver cirrhosis and Vitisin A treatment mice models were established. HE staining and Masson staining of mice liver tissue showed that disrupted hepatic architecture in the liver fibrosis group was observed and more blue collagen fiber areas. In contrast, the Vitisin A treatment group showed a reduction in liver fibrosis. Higher doses of Vitisin A showed a stronger reducing effect (Fig. 1A-B). Serum levels of ALT, AST, ALB, and TBIL were measured in four mice groups, which demonstrated dose-dependent reductions of Vitisin A in four hepatitis indicators (Fig. 1C-F). Western blot analysis revealed significant reductions in Collagen I and α-SMA protein levels by Vitisin A (Fig. 1G-I). Eventually, we observed cuproptosis level increased with increasing fibrosis in mice liver, but reversed by Vitisin A treatment in a dose- dependent manner (Fig. 1J-M). Additionally, we observed that Vitisin A significantly downregulated the expression of liver fat production markers FAS and ACC in liver tissue while enhancing the expression of β-oxidation markers CPT1 and PPARα (Fig. 1N-O). Similar results were observed in cells (Figure S1). Following Vitisin A treatment, MDA levels was significantly reduced in liver tissue (Fig. 1P).

Vitisin A inhibits liver fibrosis and cuproptosis in activated LX2s

For validation expreiments in vitro, we used TGF-β1 to activate LX2 cell line. Cell viability of LX2s was inhibited after cultured with TGF-β1 (10 ng/mL, 48 h), which is reduced by Vitisin A (Fig. 2A). As the concentration and the duration of Vitisin A treatment increased, cell viability became higher (Fig. 2B). Furthermore, we conducted cell viability assays on human hepatocytes (MIHA) and mouse hepatocytes (AML12) under conditions of liver injury and liver fibrosis induced by carbon tetrachloride (CCl₄) and transforming growth factor-β1 (TGF-β1), respectively. The results demonstrated that Vitisin A significantly restored cell viability following CCl₄ or TGF-β1 stimulation (Figure S2). Interestingly, when we cultured activated LX2 cells using ES-Cu, a cuproptosis activator, Vitisin A attenuated ES-Cu-induced cytotoxicity in activated LX2 cells (Fig. 2C). We also test expression of FDX1, a redox-active protein critical for copper-induced mitochondrial stress and other cuproptosis markers including DLAT, CTR1 and ATP7B (Fig. 2D-G). Results demonstrated that ES-Cu enhanced FDX1 and CTR1 expression while suppressing DLAT and ATP7B expression, indicating elevated cuproptosis levels that were not alleviated by Vitisin A. We separately assessed fibrosis levels and lipid synthesis and β-oxidation marker levels in LX2 cells, finding that ES-Cu eliminated Vitisin A’s suppression of cellular lipid synthesis and β-oxidation markers under hepatic fibrosis conditions while again increasing α-SMA and Collagen I expression (Fig. 2H-K). Furthermore, Vitisin A treatment significantly reduced hepatic ROS (Fig. 2L) and MDA levels (Fig. 2M), which were reversed by ES-Cu.

Vitisin A inhibits fibrosis and Cuproptosis by Nrf2/HO-1 signaling pathway

To elucidate the relationship between Vitisin A and Nrf2/HO-1 signaling pathway, we examined the expression of key proteins. Firstly, we used AutoDock Vina software for Molecular docking Analysis. The result revealed that Vitisin A binds to Nrf2 (Figure S4). Under liver fibrosis conditions, Nrf2 and HO-1 expression decreased in LX2 cells, and this decrease was restored by Vitisin A in a dose-dependent manner (Fig. 3A-C). However, Vitisin A had almost no effect on KEAP1 (Figure S3). Subsequently, we suppressed Nrf2 expression in LX2 cells using siNrf2 (Fig. 3D). Co-treatment with siNrf2 and TGF-β1 further suppressed Nrf2 and HO-1 expression, while Vitisin A partially restored their activity. Nevertheless, the combination of siNrf2 and TGF-β1 significantly promoted cuproptosis and the expression of fibrosis-related proteins (Fig. 3E-K). To elucidate the mechanism underlying Vitisin A-mediated cuproptosis inhibition, we measured CTR1 and ATP7B expression levels in hepatocytes and liver tissue. Vitisin A significantly reduced CTR1 expression while upregulating ATP7B expression (Fig. 3L). However, this protective effect was abolished in NRF2-knockout cells. ELISA results indicated that after NRF2 knockout, the effects of Vitisin A in alleviating cellular lipid synthesis and oxidative stress were significantly suppressed (Fig. 3M-N). Similar results were observed in ROS and MDA assays (Fig. 3O-P).

Vitisin A have therapeutic effect on fibrosis and cuproptosis by Nrf2/HO-1 signaling pathway in vivo

For verifying the regulatory effect of Vitisin A on the Nrf2/HO-1 signaling pathway in vivo, we used ML385 to inhibit the expression of Nrf2. Nrf2 and HO-1 were inhibited in ML385 groups and were partially recovered by Vitisin A. On the other hand, the expression of cuproptosis and fibrosis related proteins was promoted in ML385-treated groups and was inhibited in Vitisin A-treated groups (Fig. 4A-C). Furthermore, when NRF2 was downregulated in vivo, the effects of Vitisin A on lipid synthesis and oxidative stress were suppressed (Fig. 4D-F). The therapeutic effect of Vitisin A on cuproptosis-induced damage was significantly inhibited by ML385 (Fig. 4G). This inhibition was further confirmed by serum and staining analyses (Fig. 4H-I).

Discussion

In recent years, the potential of natural compounds in the treatment of liver fibrosis has attracted extensive attention22,23. Related studies have emphasized the role of flavonoids and Pyranoanthocyanin analogs in modulating inflammatory immunity24,25,26. Studies on Vitisin A have shown its anti-inflammatory and anticancer properties in diverse cellular contexts27,28. However, its efficacy and mechanism in liver fibrosis are still not well explained. How it affects cuproptosis and consequently affects other proteins and the process of hepatic fibrosis is still not well understood. Our results showed that Vitisin A significantly inhibited cuproptosis and the progression of hepatic fibrosis, as well as upregulated Nrf2/HO-1 while suppressed the expression of copper apoptosis-associated proteins. Vitisin A has been shown to be involved in the regulation of multiple proteins, contributing to its role in antifibrotic effects. Although further validation of its effects in clinical trials is required, the role of Vitisin A in attenuating hepatic fibrosis has been demonstrated in vivo and vitro.

Cuproptosis has been shown to be closely associated with inflammatory responses and metabolic mechanisms in liver fibrosis29,30. Excessive copper ions bind to the lipoylated enzymes in the tricarboxylic acid cycle, which leads to lipoylated protein aggregation, proteotoxic stress, and ultimately cell death31,32. Tian et al. confirmed that Diallyl trisulfides (DATs) induces phase separation of RAB18 as well as activating cuproptosis to selectively protect hepatocytes33. It is evident that copper death and liver fibrosis is not a simple positive linear relationship, which involves complex regulatory processes of copper metabolism. Our results also indicate that ES-Cu elevated the expression of DLAT in liver fibroblasts, while the DLAT expression is rather decreased when acting simultaneously with Vitisin A, which suggesting direct effects of copper ion concentration on hepatic fibrotic cells. Our experiments also showed that Vitisin A reduced hepatic fibrosis-associated proteins as well as cuproptosis markers in a dose-dependent and time-dependent manner. Notably, Vitisin A did not reverse hepatic fibrosis, but demonstrated a role in delaying the progression of hepatic fibrosis. The effects of Vitisin A were not compared with other drugs, and further studies are needed to determine whether it may have synergistic effects with other drugs. In addition, Vitisin A may attenuate fibrosis through crosstalk between cuproptosis and metabolic pathways, though further mechanistic validation is required.

The Keap1/Nrf2/HO-1 pathway regulates oxidative stress and inflammatory responses, with component expression levels inversely correlated with fibrosis severity34,35,36. Keap1 inhibits the activity of Nrf2 by promoting its ubiquitination degradation, thereby promoting the progression of liver fibrosis37,38,39. In addition, post-translational modifications of Keap1 (e.g., cysteine oxidation) could indirectly regulate Nrf2 activity, thereby modulating liver fibrosis40,41. Researchers have proved the correlation between cuproptosis and Keap1/Nrf2/Atp7b pathway15. Our experimental results showed that Vitisin A promoted Nrf2 signaling and its downstream targets, but did not affect the expression level of Keap1, suggesting that Vitisin A is an upstream regulator of Nrf2. Further experiment of Nrf2 inhibition demonstrated molecular mechanisms of Vitisin A action. Vitisin A may disrupt Keap1-Nrf2 binding, preventing Nrf2 ubiquitination and degradation, which needs more experimental evidence.

Conclusion

In conclusion, our present study demonstrated that Vitisin A inhibited hepatic stellate cell activation and hepatic fibrosis in mice in a concentration-dependent manner. Nrf2 knockdown via siRNA and inhibition by ML 385 abolished Vitisin A’s anti-fibrotic effects both in vivo and vitro. ES-Cu attenuated Vitisin A-mediated fibrosis suppression, which revealed a molecular relationship between Vitisin A and cuproptosis. The inhibition of Nrf2 by siRNA also blocked Vitisin A inhibiting liver fibrosis, which further suggests that Vitisin A is an upstream regulator of Nrf2. Vitisin A may bind to Nrf2 and upregulating degradation of its ubiquitination level, although post-translational modification studies are required to validate. Combined with results from in vivo and in vitro trials, Vitisin A demonstrates clinical therapeutic potential for the treatment of liver fibrosis.