Hepatocyte growth factor alleviates alcoholic cardiomyopathy through the Nrf2 pathway—a focus on iron metabolism

Introduction

The issue of alcohol abuse has been a global health concern for a long time. Excess or harmful alcohol use contributes significantly to the disease burden due to mental health problems or non-mental diseases such as cardiac diseases. A previous study suggests that men who consume moderate-to-large amounts of alcohol regularly might develop atrial fibrillation (AF), and over-consumption of alcohol leads to alcoholic cardiomyopathy (ACM), which manifests as myocardial injury, ventricle dilation and cardiac dysfunction (1). The main active ingredient of various alcoholic beverages is ethanol, of which its primary metabolite produced by the action of ethanol dehydrogenase in the body, can be a potential killer of cardiovascular systems. In recent years, an increasing number of studies have confirmed that acetaldehyde is not only the main culprit in inducing various types of cancer after drinking alcohol, but also a major toxic substance that causes alcoholic heart disease (2).

Numerous novel cell death modes, including necroptosis, pyroptosis, autophagy, and ferroptosis, have been discovered in recent research that have disproved earlier theories regarding cell death (necrosis and apoptosis), and they have contributed significantly to our understanding of cardiovascular disorders. In 2012, the term ferroptosis was first described as a distinct iron-dependent form of programmed cell death (3). While the induction of ferroptosis is linked to various diseases (4), a connection between ferroptosis and ACM has yet to be proven.

Hepatocyte growth factor (HGF), also known as scatter factor, is necessary for embryogenesis; and HGF/c-Met has complex functions in cellular proliferation, survival, angiogenesis, and tissue repair (5). Hepatocyte or liver models have been the major focus of recent HGF-related investigations on alcohol-induced injury. Nevertheless, the reaction between HGF and alcoholic cardiac intoxication is still unknown. In the current work, a model of alcoholic myocardial damage was created, and it was predicted and validated that HGF protects the myocardium via modulating ferroptosis. We present this article in accordance with the MDAR and ARRIVE reporting checklists (available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-1111/rc).

Methods

Reagent and antibodies

Recombinant rat HGF protein (P1779) was obtained from FineTest (Wuhan, China); rabbit polyclonal anti-rat Fpn1 (26601-1-AP), rabbit polyclonal anti-rat DMT1 (20507-1-AP), goat anti-mouse or anti-rabbit secondary antibodies and mouse monoclonal anti-β-actin from Proteintech (Wuhan, China); rabbit polyclonal anti-rat ferritin heavy chain (FTH1) (PA5-27500) from Thermofisher (Waltham, USA). The Bicinchoninic Acid protein assay kit (71285-M), calcein-AM (206700) and 3-hydroxy-1,2-dimethyl-4(1H)-pyridone (379409) from Sigma-Aldrich (St. Louis, USA); the Trizol reagent, RIPA protein lysis buffer, malondialdehyde (MDA) kit (S0131S), Cell Counting Kit-8 (CCK8), calcein staining kit (C2013S) and hematoxylin-eosin (HE) staining kit (C0105S) from Beyotime (Shanghai, China); the glutathione peroxidase 4 (GPX4) (CSB-EL009869RA) and HGF (CSB-E07346r) enzyme-linked immunosorbent assay (ELISA) kit from Cusabio (Wuhan, China). All animal experiments were performed under a project license (No. IACUC-2111096) granted by Ethics Committee of Nanjing Medical University, in compliance with the institutional guidelines for the care and use of animals. A protocol was prepared before the study without registration.

Establishment of ethanol-induced cardiomyocyte injury

Dulbecco’s modified Eagle’s medium (DMEM, Gibco, Waltham, USA) was used to cultivate heart-derived H9c2 cells (Chinese Academy of Science Cell Bank, Shanghai, China) at 37 ℃ along with 10% fetal bovine serum and 1% penicillin-streptomycin solution in humidified air (5% CO₂). A minimal essential serum-free medium was used for 12 hours prior to treatments, and the medium was switched after 24 hours. Depending on the rate of cell development, it was switched again for 1–2 days. The cells were monitored each day. Cardiomyocytes were given two phosphate buffer saline (PBS) rinses before being put to the medium with the serum removed and being treated with 1% ethanol for 24 hours. After fluid exchange, an effective in vitro model of alcohol-induced cardiac damage was developed and utilized for further research.

Preparation of ACM model in rats

Twenty-four male Sprague Dawley rats with a body weight of 250–350 g were selected and randomly allocated into model group and control (ctrl) group. The sample size was calculated by referring to previous similar studies and taking into account statistical power. For model preparation, in the first week, the rats were allowed to drink liquor containing 10% alcohol (prepared by drinking water) at will, and gavage the liquor containing 60% alcohol (10 mL/kg) once a day. In the second week, rats could drink 20% alcohol-containing liquor every day, and gavage twice a day with 60% alcohol-containing liquor (15 mL/kg). From the 3rd week to the 13th week, rats were given 30% alcohol-containing liquor every day, and gavage 60% alcohol liquor three times a day (20 mL/kg). Finally, 10 rats survived in each of the model group (2 died and excluded) and the control group (2 died and excluded) for subsequent experiments. Each rat was regarded as an independent experimental unit. In the experiment, inbred rats were selected to reduce individual genetic variations. Meanwhile, rats of the same strain, age at weeks and gender were used to avoid deviations caused by species differences. In addition, the weight range and feeding methods were unified to further reduce deviation.

Tissue staining

After anesthetization, rats’ heart tissues were extracted by thoracotomy and encased in paraffin were cut into 4-µm sections, dewaxed with xylene, and then rehydrated using a serial alcohol gradient. Following a 1× PBS wash, the slides were dehydrated using escalating ethanol and xylene concentrations and stained with hematoxylin and eosin.

ELISA assay

HGF and the GPX4 activity were measured using corresponding Elisa kits as instructed. Prepare reagents, standards, and samples as instructed. Add 100 µL of standard/sample per well, incubate at 37 ℃ for 2 hours. Aspirate without washing, then add 100 µL Biotin-antibody (1×) and incubate at 37 ℃ for 1 hour. Wash wells 3× with 200 µL Wash Buffer. Add 100 µL HRP-avidin (1×), incubate at 37 ℃ for 1 hour, followed by 5× washes. Add 90 µL TMB Substrate (protect from light), incubate for 15 minutes. Stop the reaction with 50 µL Stop Solution. Measure optical density (OD) at 450 nm, using 540/570 nm for correction.

MDA assay

The cells were plated at a density of 2.0×10⁵ cells/well in 12-well plates, and after exposure as anticipated, they were harvested by trypsinization, cellular extracts were made using cell lysis buffer, and the lysed cells were centrifuged at 1,600 rpm for 10 minutes to remove debris. The protein content and MDA level were measured in the supernatant, and each test tube was filled with 100 µL cell homogenates, 50 µL 0.37% thiobarbituric acid (TBA), and 150 µL diluent of TBA. A hot water bath was used to incubate the mixture for fifteen minutes. The reaction mixture was centrifuged at 1,000 rpm for 10 minutes after it had cooled to room temperature. A 96-well plate was then filled with 200 µL of supernatant, and the absorbance was measured at 540 nm using a microplate reader. Control was set at 490 nm. To measure the protein concentration, an improved BCA protein assay kit was employed.

Western blot

A protein extraction kit was used to extract the proteins from each group of cells. Before electrophoresis, 20 µL of the samples were loaded on a 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), then transferred to a polyvinylidene fluoride (PVDF) membrane, blocked with 5% skim milk, and incubated with the corresponding primary antibody for an overnight period at 4 ℃. After three washes, add the matching secondary antibody, let the mixture sit at room temperature for one hour, and then use the ECL kit to image, detect, and analyze the protein.

Real-time polymerase chain reaction (PCR)

Samples were processed for total RNA extraction in accordance with the guidelines for the TriFast^TM Purification Kits. Using the QIAGEN One Step RT-PCR Kit, extracted RNA was transformed into cDNA. The primers used were as follows: hepcidin, sense primer 5'-GAAGGCAAGATGGCACTAAGCA-3', antisense primer 5'-TCTCGTCTGTTGCCGGAGATAG-3'.

Assessment of intracellular labile iron pool (LIP)

Using the fluorescent iron sensor calcein-AM, flow cytometry was used to measure the intracellular labile iron content (6). Cells were treated for 30 minutes at 37 ℃ in the dark with 0.25 µM calcein-AM. After removing the excess calcein-AM with two PBS washes, cells were either left untreated or treated with 200 µM of the iron chelator L1 [3-hydroxy-1,2-dimethyl-4(1H)-pyridone]. The FACS BD LSRFortessa^TM X-20 cytofluorometer was used to conduct the analysis. The LIP was represented by the variation in cellular fluorescence before and after incubation with L1.

CCK8 test and calcein-AM staining for cell viability determination

Prepare 100 µL of cell suspension on 96-well plates, corresponding to different interventions, according to the CCK8 kit instructions. The plates were pre-incubated in an incubator at 37 ℃ and 5% CO₂ for 24 hours. Incubate each well again for 3 hours at 37 ℃ with 5% CO₂ after adding 10 µL of the CCK8 solution. The absorbance at 450 nm was measured.

Ethanol-treated cells were co-stained with calcein-AM for 0.5 h, then washed with fresh medium two times and imaged by a fluorescence inverted microscope. Calcein-AM exhibited no fluorescence. After entering the H9c2 cells, it was hydrolyzed by endogenous esterase to generate calcein, a polar molecule with a significant negative charge that cannot pass through the cell membrane and was kept in the cell, emitting bright green fluorescence. Compared with other probes of the same kind, calcein-AM is one of the most ideal fluorescent probes for staining live cells because it has very low cytotoxicity, hardly affects cell functions such as cell proliferation or lymphocyte chemotaxis, and is less sensitive to pH value.

Target prediction

The ferroptosis database (www.zhounan.org/ferrdb) was searched, and the driver, marker, suppressor, and unclassified datasets were pooled to form the ferroptosis-related gene ensemble, which yielded 564 items after de-duplication. The GeneCards disease database (https://www.genecards.org/) was also searched using the keyword ACM, and the disease-related gene dataset was obtained, screening 372 genes with a relevance score of 10 or higher. The obtained target intersections were imported into the STRING database (https://cn.string-db.org/) for analysis, isolated nodes were removed, and the analysis results were imported into Cytoscape software to create network diagrams, while network parameters were addressed using the ClusterViz application’s MCODE algorithm.

Gene-regulating experiment

Fluorescent-labeled adenovirus packaged with Nfe2l2 (Nfe2l2-siRNA) was designed and synthesized by GeneChem (Shanghai, China) (7). Then, cells were infected with Nfe2l2-siRNA and scramble at a multiplicity of infection (moi) of 100 for 48 h.

Statistical analysis

Each lab experiment was replicated three times. Data were presented as mean ± standard deviation (SD). All statistical calculations were made using SPSS software, version 19.0. To compare group differences, a one-way analysis of variance and an unpaired t-test were utilized. Statistically significant was determined as P<0.05.

Results

Establishment of alcohol-injured rats with cardiomyopathy

The effectiveness of the modeling was assessed using two-dimensional echocardiography and HE staining of the myocardium after rats were raised on a fixed formulation of alcohol on a regular schedule. Both the fractional shortening (FS) and the ejection fraction (EF) indicate the functioning state of the heart. Rats fed alcohol over a prolonged period of time had a substantial decline in cardiac function [Ctrl group (n=10) vs. ACM group (n=10), EF: 72.41%±3.78% vs. 65.14%±2.94%, FS: 43.40%±3.51% vs. 37.21%±2.26%] (Figure 1A). The thickness of the left ventricular wall is reflected by the left ventricular anterior wall end-systolic dimensions (LVAWs) and left ventricular posterior wall end-systolic dimensions (LVPWs). Rats fed alcohol for a prolonged period of time had thinner ventricular walls (Ctrl group vs. ACM group, LVAWs: 3.13±0.05 vs. 3.01±0.09 mm, LVPWs: 3.22±0.06 vs. 3.13±0.07 mm) (Figure 1B). Stroke volume, another marker of heart condition, was considerably lower in the ACM group (Ctrl group vs. ACM group, 275.43±12.86 vs. 255.01±10.75 µL) (Figure 1C). Rats in the ACM group had atrophy and irregular fractures in their cardiac muscle fibers, according to HE staining, whereas the cardiomyocytes in the Ctrl group were stuffed with healthy myocardial cells, and the myocardial fibers were neatly aligned (Figure 1D). Images of the heart’s function under M-mode cardiac ultrasonography are also displayed in Figure 1D.

Figure 1 Altered cardiac structure and function in ACM rats. (A) EF and FS reduction after alcohol induction; (B) ventricle wall thickness decline in alcohol fed rats; (C) stroke volume drop in the established ACM rats; (D) M-mode echocardiography displaying cardiac functional changes, and H&E staining showing altered cardiac anatomy. **, P<0.01. ACM, alcoholic cardiomyopathy; Ctrl, control; EF, ejection fraction; FS, fractional shortening; H&E, hematoxylin and eosin; LVAWs, left ventricular anterior wall end-systolic dimensions; LVPWs, left ventricular posterior wall end-systolic dimensions; M-mode Echo, M-mode echocardiography.

Serum HGF levels

The serum HGF levels were evaluated by ELISA. Comparing the experimental group to the control group, the blood levels of HGF tended to rise following alcohol induction (Ctrl group vs. ACM group, 4.84±0.94 vs. 6.50±1.07 ng/mL) as depicted by the violin map with all points in Figure 2A.

Figure 2 Levels of serum HGF and effects of ethanol on cardiomyocytes. (A) Violin map displaying HGF concentrations; (B) CCK8 test for cardiomyocyte viability in response to various strategies for alcohol concentration; (C) effects of various alcohol intervention doses on cell survival by calcein staining. **, P<0.01. ACM, alcoholic cardiomyopathy; CCK8, Cell Counting Kit-8; Ctrl, control; HGF, hepatocyte growth factor.

Effects of ethanol on cardiomyocytes

Figure 2B illustrates the detrimental effects of different ethanol concentrations on cardiomyocyte viability by CCK8 assay. The 200 mM ethanol appeared to slightly diminish cardiomyocyte survival. As demonstrated in Figure 2C, calcein staining revealed that the majority of the visual field was still filled with live cells. When the concentration of ethanol increased to 400 mM, the survival rate of cells decreased sharply, and calcein staining showed that there were still many surviving cells. Higher ethanol concentrations would cause a further decrease in cell survival rate, but the absolute number of surviving cells was already fairly small and scattered across the calcein staining region, making it unsuitable for further investigation.

Effect of exogenous HGF on ferroptosis in cardiomyocytes

When 400 mM ethanol was added to normal cardiomyocytes, cell survival decreased to 48.67%. However, HGF pretreatment of cardiomyocytes prior to ethanol exposure was found to significantly improve cell viability (86.99%) as illustrated in Figure 3A. This experiment was carried out independently for the GPX4 (Figure 3B) and LIP (Figure 3C) assay to determine whether HGF has an impact on ferroptosis. Figure 3B,3C demonstrates that the addition of HGF reversed the ethanol-induced reduction in GPX4 and promotion in LIP in injured cardiomyocytes. In contrast, the administration of Fer-1, a specific inhibitor of ferroptosis, had effects that were similar to those of HGF, indicating that both HGF and Fer-1 contributed to reversing ferroptosis. Figure 3D visualizes cell survival in each group under different interventions by calcein staining. Cardiomyocytes were treated for 1 hour with various doses of HGF (0, 0.2, 0.6, 0.8, or 1.0 nM), and subsequently given 24 hours of treatment with 400 mM ethanol. Each group’s MDA (Figure 3E) and GPX4 (Figure 3F) concentrations, along with LIP levels (Figure 3G), were then tested. Following ethanol stimulation, the HGF concentration that was both acceptable and efficient is 0.6 nM, since GPX4 had the maximum value and MDA represented the smallest value. At this time, LIP levels were the lowest across the groups. This implies that HGF values of 0.6 nM correspond to the lowest levels of ferroptosis in cardiomyocytes. Figure 3H-3J demonstrates that knockdown of Nfe2l2 switched the HGF-induced elevation in GPX4, together with a decline in MDA and LIP in ethanol-injured cardiomyocytes. Figure 3K displays that HGF preconditioning enhanced the quantity of cells that survived alcohol treatment, whereas Nfe2l2 knockdown dramatically decreased HGF’s protective impact.

Figure 3 Effects of exogenous HGF on ferroptosis in cardiomyocytes. (A) The effect of HGF on cell viability (CCK8); (B) the effect of HGF on GPX4; (C) the effect of HGF on LIP; (D) the effect of HGF on cell survival by calcein staining; (E) MDA concentrations of different HGF doses; (F) GPX4 concentrations of different HGF doses; (G) LIP levels of different HGF doses; (H) MDA concentrations following Nfe2l2 knockdown in ethanol-treated cells; (I) GPX4 concentrations following Nfe2l2 knockdown in ethanol-treated cells; (J) LIP levels following Nfe2l2 knockdown in ethanol-treated cells; (K) Cell survivals by calcein staining following Nfe2l2 knockdown in ethanol-treated cells. **, P<0.01; ns, P>0.05. CCK8, Cell Counting Kit-8; GPX4, glutathione peroxidase 4; HGF, hepatocyte growth factor; LIP, labile iron pool; MDA, malondialdehyde; MFI, mean fluorescence intensity; PBS, phosphate buffered saline; scramble, randomly interrupted sequence; siRNA, small interfering RNA.

Key target of HGF protection

An inquiry and analysis using public databases was conducted to find additional potential targets for HGF’s cardioprotective actions. Forty-eight genes were related to the two datasets, which were taken from the GeneCards and FerrDb databases, respectively (Figure 4A). After removing isolated nodes and duplicate values, there were 45 items. Two sub-networks were identified using ClusterViz’s MCODE technique, the major of which includes 26 significant nodes and 294 edges with an average score of 23.52. The parameters of the chosen sub-network are displayed in Table 1 in descending order of score. We ultimately discovered Nfe2l2 to be an essential target (Figure 4B). Nuclear factor erythroid 2-related factor 2 (Nrf2) is a protein encoded by the Nfe2l2 gene. Western blot assay showed that cardiomyocytes exposed to HGF for 30 minutes expressed more Nrf2 in both the cytoplasm and the nucleus, with the proportion of Nrf2 in the nucleus rising (Figure 4C). Next, we utilized small interfering RNA to silence Nrf2 expression, verifying the key role of Nrf2 in ferroptosis, as displayed in Figure 3. Research on ethanol-exposed cardiomyocytes revealed that exogenous HGF drastically restricted MDA and LIP levels while substantially drove GPX4 activity and cell survival. However, silencing Nfe2l2 resulted in rises in MDA and LIP along with declines in GPX4 and active cell quantity.

Figure 4 Investigating the key node of HGF against ferroptosis in cardiomyocytes. (A) The intersection of two datasets; (B) subnetwork analysis and critical target mining; (C) Nrf2 expression in the cytoplasm and nucleus following HGF treatment. HGF, hepatocyte growth factor.

Effect of HGF on proteins involved in iron metabolism

Hepcidin, a hormone secreted by the liver that controls iron by interacting with its receptor ferroportin (Fpn), is known to influence systemic iron distribution and storage. To that purpose, this work investigated alterations in the hepcidin/Fpn1 pathway with or without HGF pretreatments, in ethanol influenced cells or not respectively (Figure 5A). In cardiomyocytes that had not been affected by ethanol, little difference was found after HGF intervention in the levels of hepcidin mRNA expression; however, in the ethanol-induced cardiomyocyte, hepcidin mRNA levels were significantly lower in the HGF pretreatments group compared to those without HGF administration. At the same time, hepcidin expression was stronger in the ethanol-stimulated group compared to normal cardiomyocytes at baseline. To explore the effect of HGF on Fpn1 and whether it acts through the crucial node of Nrf2, gene knockdown experiment and western blot assays were performed (Figure 5B,5C). HGF pretreatment significantly boosted the expression of the Fpn1 protein in ethanol-induced cardiomyocytes. However, Fpn1 protein was reduced significantly as a result of Nfe2l2 knockdown. DMT1, namely divalent metal-ion transporter 1, is extensively expressed in mammals and is involved in the transport of various metal ions. DMT1 contributes specifically to total iron flux. HGF treatment enhanced the expression of DMT1 protein in ethanol-pretreated cells, while decreasing Nfe2l2 expression at the same time, had no further effects on DMT1 expression. FTH1 is also extremely sensitive to HGF intervention. Contrary to the above-mentioned two iron metabolism-related proteins, FTH1 protein expression dramatically decreased after HGF administration into the ethanol group, and Nfe2l2 suppression switched the decline tendency of FTH1 levels.

Figure 5 Effects of HGF on proteins involved in iron metabolism. (A) Hepcidin mRNA expression was raised by ethanol stimulation and decreased by HGF treatment; (B,C) alterations in Fpn1, DMT1 and FTH1 caused by Nfe2l2 knockdown after HGF pretreatment detected by western blotting under ethanol exposure. **, P<0.01; ns, P>0.05. DMT1, divalent metal-ion transporter 1; Fpn1, ferroportin; FTH1, ferritin heavy chain 1; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; HGF, hepatocyte growth factor; PBS, phosphate buffered saline; scramble, randomly interrupted sequence; siRNA, small interfering RNA.

Discussion

Numerous studies have shown that HGF can be utilized as a marker for cardiovascular conditions since different cardiovascular problems occur with fluctuating HGF levels. Previous evidence has revealed that the endogenous HGF expression levels may briefly increase to a peak in the early stages of disease before gradually declining (8,9). As a result, increased levels of circulating HGF are indicative of the early stages of some cardiac diseases. Conditions including heart failure (10), acute myocardial infarction (11), hypertension (12), and atherosclerosis (13) are all associated with higher plasma HGF concentrations.

Alcohol induces oxidative stress, which has a deleterious influence on the heart in vivo, with lipid peroxidation being a major sign (2). When the ethanol concentration reached 400 mM, it can have both negative effects and preserve enough cardiomyocytes for additional research. Pretreatment of cardiomyocytes with HGF prior to ethanol exposure was found to significantly enhance cell viability. Our studies demonstrated a connection between ethanol-induced cardiomyocyte damage and anomalies in iron metabolism in cardiomyocytes. The iron-dependence and lipid peroxidation-dependence are the two most crucial elements of ferroptosis (14). Although HGF exhibits anti-oxidative stress properties, its connection to ferroptosis is not well understood. Pretreating ethanol-exposed cardiomyocytes with a range of HGF doses in a stepwise manner allowed for the determination of the most suitable and efficient concentration of HGF intervention. The degree of lipid peroxidation was then estimated by measuring the amounts of MDA and GPX4, and the degree of iron metabolism inhomoeostasis was assessed by examining the level of LIP. In a model of ethanol-induced cardiomyocyte damage, it was later discovered that the effects of HGF were comparable to those of Fer-1, a particular inhibitor of iron metabolism. The data supporting the aforementioned assertions showed that HGF did affect ferroptosis and that this effect was similar to that of the protective effect from Fer-1.

The current study’s discovery that HGF reduces ferroptosis in cardiomyocytes raises the fascinating question of how HGF interacts with important nodes of the iron metabolism network. Iron intake, iron storage, and iron excretion are the three primary roles of the complex network that controls the body’s iron metabolism (15). The hepatocyte-derived protein hepcidin modulates iron metabolism systemically and is a major regulator of iron homeostasis (16). A previous study found that HGF inhibits the expression of hepcidin mRNA, and that this effect is reversed by blocking the met receptor (17). Hepcidin regulates dietary iron absorption by suppressing DMT1 and Fpn1 apical transporter expression in the duodenum (18). Under physiological conditions, Fpn1 functions as the body’s only iron exporter (19), and the amount of DMT1 shows how actively divalent iron ions are transported inside cells (20). In a recent study, Qin’s group discovered that pharmacological therapies that increase hepcidin expression are linked to decreased expression of Fpn1 and DMT1 (21). The majority of our results are consistent with the earlier findings. HGF overexpression was linked to a significant increase in the protein production of the Fpn1 and DMT1 in ethanol-injured cardiomyocytes. By binding iron ions, a protein known as ferritin acts as the body’s ferrous storage (22). The ferritin heavy chain predominates in the heart, whereas light chain-rich ferritin is more common in iron-storing organs like the liver (23). In this investigation, HGF upregulation decreased FTH1 expression in the ethanol-induced cardiomyocytes. Briefly, HGF showed increased iron absorption and iron excretion from cardiomyocytes, as well as decreased iron storage, in the ethanol-injured condition. The cumulative consequence of HGF action on these important iron metabolism nodes is, in fact, ferroptosis suppression. It is reasonable to assume that in this instance, HGF’s stimulation of iron excretion greatly surpasses its impact on iron intake and storage, ultimately resulting in a decrease in cellular iron.

Following an investigation of public databases to identify the intersection of genes associated with ferroptosis and ACM, a core regulatory network was built to identify putative genes with greater weights. Nfe2l2 may be a significant factor influencing iron metabolism and oxidative stress, according to the findings. Inhibitors that specifically target one ferroptosis-inducing enzyme or toxic ferroptosis byproducts, however, might not be as effective in blocking ferroptosis as those that directly affect Nrf2, which acts on several ferroptosis-actors upstream (24). In actuality, Nrf2 is located at the junction of lipid, redox, and iron homeostasis, which forces us to determine its precise function in ferroptosis (25). It has been reported that Nrf2 negatively regulates ferroptosis (26), and ferroptosis is apparently suppressed by Nrf2 stability (27). A subsequent experiment revealed that once HGF was added, ethanol-exposed cardiomyocytes’ LIP intensity decreased; nevertheless, Nrf2 silencing reversed this effect. In addition, the impacts of Nrf2 regulation on lipid peroxidation exhibited a pattern of tendency similar to that of LIP. Since it is now known that HGF may affect cardiomyocyte ferroptosis through Nrf2, the question of what function Nrf2 plays in the network of HGF-regulated cardiac iron metabolism then comes up. In a cellular model of ethanol-induced heart damage, overexpression of HGF and concomitant Nfe2l2 silencing seems to have a number of distinct alternative effects on important proteins of the iron metabolism network. Fpn1 expression was downregulated and FTH1 expression was increased when Nfe2l2 was knocked down because Nfe2l2 silencing stopped the HGF-induced increase in Fpn1 expression and the decrease in FTH1 expression. Nrf2 has been identified to regulate a number of genes (FTL/FTH1, FPN1, and GPX4) involved in iron export and storage (28-30). This is most likely because Nrf2 and BACH1 influence ferroportin (FPN1), ferritin’s heavy and light chains, and ferritin’s ARE sequence during erythropoiesis (31). Ferritin helps to maintain the homeostasis of the iron pool by storing excess iron in conjunction with FTH1 and ferritin light chain (FTL) (32-33). However, it has currently been proven that Nrf2 controls ferroptosis through modulating FTH1, but not FTL or TFR1 (34). FTH1 protein levels are more dependent on Nrf2-mediated transcription than FTL (35). In addition, it is the fact that organs with rapid iron turnover, like the heart, have more FTH1 because it has ferroxidase activity, whereas organs that store iron, like the liver, have more FTL because it makes it easier to store iron in the core (36). In other words, the ferritin heavy chain is the predominant component in the heart, as opposed to iron-storage organs like the liver where light chain-rich ferritin is more abundant (23). For this reason, in order to better understand how Nrf2-regulated iron storage works, the current investigation set FTH1 as the primary research target. In the subsequent experiment, this study was able to demonstrate that while HGF may increase DMT1 expression, additional DMT1 alterations were not significantly impacted by Nfe2l2 silencing. DMT1 gene expression was not correlated with Nrf2 in cardiac tissues, but it was in hepatic tissues, which may be because iron content in the heart controls DMT1 expression more so than oxidative stress does in the liver (37). This implies that even while Nrf2 is a key target of HGF to stop ferroptosis, it also plays a role in the iron metabolism regulating system. It is not, however, the sole HGF hub that has anti-ferroptosis properties. The specific phenomenon for DMT1 indicates another factor-mediated mechanism (except Nrf2) between HGF and iron metabolism.

Standard ACM management focuses on complete abstinence alongside guideline-directed therapy for heart failure with reduced ejection fraction (HFrEF) [angiotensin-converting enzyme inhibitor (ACEi)/angiotensin-receptor blocker (ARB)/angiotensin receptor-neprilysin inhibitor (ARNI), beta-blocker, mineralocorticoid receptor antagonist (MRA), sodium-glucose cotransporter-2 inhibitor (SGLT2i)], and device therapy if warranted. HGF may serve as an adjunctive disease-modifying agent that enhances, rather than supplants, guideline-directed medical therapy (GDMT). HGF’s protective mechanisms focus on pathways inadequately addressed by GDMT, allowing for synergy with ARNI, SGLT2i, and MRA. From a biological standpoint, the efficacy of HGF is compelling, and there exists substantial preclinical research data to substantiate its cardioprotective properties. However, there is currently a dearth of direct clinical trial data; as a result, properly planned, mechanism-oriented early trials are necessary to promote its implementation. Limitations of the study: this study did not explore the changes and interactions of other pathways in ACM. Moreover, no further in-depth research was conducted on the long-term safety of HGF administration.

Conclusions

Altogether, our results show that HGF possesses cardioprotective qualities that may prevent injury from alcohol. HGF acts as an anti-ferroptosis agent by increasing Nrf2 expression levels, which reduces lipid peroxidation and the LIP. Nevertheless, the way that HGF regulates iron metabolism is highly intricate, and since Nrf2 controls a portion of the iron metabolism node, more investigation is still required to identify a more complete and specific regulatory mechanism.

Acknowledgments

Prof. Wei Sun, Prof. Xiangqing Kong and Prof. Peng Li provided support for the effective execution of this study by helping to complete the work and enable the execution of the experiment. We would like to extend our heartfelt thanks to them for their assistance in these endeavors.

Footnote

Reporting Checklist: The authors have completed the MDAR and ARRIVE reporting checklists. Available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-1111/rc

Data Sharing Statement: Available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-1111/dss

Peer Review File: Available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-1111/prf

Funding: This work was supported by National Natural Science Foundation of China (No. 81900337, to J.Z.).

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-1111/coif). J.Z. reports funding from National Natural Science Foundation of China (No. 81900337). The other authors have no conflicts of interest to declare.

Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. All animal experiments were performed under a project license (No. IACUC-2111096) granted by Ethics Committee of Nanjing Medical University, in compliance with the institutional guidelines for the care and use of animals.

Open Access Statement: This is an Open Access article distributed in accordance with the Creative Commons Attribution-NonCommercial-NoDerivs 4.0 International License (CC BY-NC-ND 4.0), which permits the non-commercial replication and distribution of the article with the strict proviso that no changes or edits are made and the original work is properly cited (including links to both the formal publication through the relevant DOI and the license). See: https://creativecommons.org/licenses/by-nc-nd/4.0/.

References

1. Lee DI, Kim S, Kang DO. Exploring the complex interplay between alcohol consumption and cardiovascular health: Mechanisms, evidence, and future directions. Trends Cardiovasc Med 2025;35:243-53. [Crossref] [PubMed]
2. Lu Q, Qin X, Chen C, et al. Elevated levels of alcohol dehydrogenase aggravate ethanol-evoked cardiac remodeling and contractile anomalies through FKBP5-yap-mediated regulation of ferroptosis and ER stress. Life Sci 2024;343:122508. [Crossref] [PubMed]
3. Dixon SJ, Lemberg KM, Lamprecht MR, et al. Ferroptosis: an iron-dependent form of nonapoptotic cell death. Cell 2012;149:1060-72. [Crossref] [PubMed]
4. Berndt C, Alborzinia H, Amen VS, et al. Ferroptosis in health and disease. Redox Biol 2024;75:103211. [Crossref] [PubMed]
5. Gao J, Wang X, Zhang L, et al. Unveiling the role of c-Met: A promising target for cardiovascular disease. Pharmacol Res 2025;219:107893. [Crossref] [PubMed]
6. Battaglia AM, Sacco A, Vecchio E, et al. Iron affects the sphere-forming ability of ovarian cancer cells in non-adherent culture conditions. Front Cell Dev Biol 2023;11:1272667. [Crossref] [PubMed]
7. Gu H, Zhu Y, Zhou Y, et al. LncRNA MALAT1 Affects Mycoplasma pneumoniae Pneumonia via NF-κB Regulation. Front Cell Dev Biol 2020;8:563693. [Crossref] [PubMed]
8. Pannitteri G, Petrucci E, Testa U. Coordinate release of angiogenic growth factors after acute myocardial infarction: evidence of a two-wave production. J Cardiovasc Med (Hagerstown) 2006;7:872-9. [Crossref] [PubMed]
9. Suzuki H, Murakami M, Shoji M, et al. Hepatocyte growth factor and vascular endothelial growth factor in ischaemic heart disease. Coron Artery Dis 2003;14:301-7. [Crossref] [PubMed]
10. Chen HL, Zhu XT, Zhang W, et al. Hepatocyte growth factor and B-type natriuretic peptide as independent predictors of mortality in HFpEF patients. Front Cardiovasc Med 2025;12:1512411. [Crossref] [PubMed]
11. McQuaig R, Dixit P, Yamauchi A, et al. Combination of Cardiac Progenitor Cells From the Right Atrium and Left Ventricle Exhibits Synergistic Paracrine Effects In Vitro. Cell Transplant 2020;29:963689720972328. [Crossref] [PubMed]
12. Otsuka M, Adachi H, Jacobs DR Jr, et al. Serum hepatocyte growth factor and cancer mortality in an apparently healthy Japanese population. J Epidemiol 2012;22:395-401. [Crossref] [PubMed]
13. Sommer P, Schreinlechner M, Noflatscher M, et al. Hepatocyte growth factor as indicator for subclinical atherosclerosis. Vasa 2024;53:120-8. [Crossref] [PubMed]
14. Ye Z, Liu W, Zhuo Q, et al. Ferroptosis: Final destination for cancer? Cell Prolif 2020;53:e12761. [Crossref] [PubMed]
15. Yousefi MH, Masoudi A, Saberi Rounkian M, et al. An overview of the current evidences on the role of iron in colorectal cancer: a review. Front Oncol 2025;15:1499094. [Crossref] [PubMed]
16. Liu J, Zhao J, He J, et al. Hepcidin mediates the disorder of iron homeostasis and mitochondrial function in mice under hypobaric hypoxia exposure. Apoptosis 2025;30:1076-91. [Crossref] [PubMed]
17. Goodnough JB, Ramos E, Nemeth E, et al. Inhibition of hepcidin transcription by growth factors. Hepatology 2012;56:291-9. [Crossref] [PubMed]
18. Saad HKM, Taib WRW, Ismail I, et al. Reduced hepcidin expression enhances iron overload in patients with HbE/β-thalassemia: A comparative cross-sectional study. Exp Ther Med 2021;22:1402. [Crossref] [PubMed]
19. Steinbicker AU, Bartnikas TB, Lohmeyer LK, et al. Perturbation of hepcidin expression by BMP type I receptor deletion induces iron overload in mice. Blood 2011;118:4224-30. [Crossref] [PubMed]
20. Tian J, Zheng W, Li XL, et al. Lower Expression of Ndfip1 Is Associated With Alzheimer Disease Pathogenesis Through Decreasing DMT1 Degradation and Increasing Iron Influx. Front Aging Neurosci 2018;10:165. [Crossref] [PubMed]
21. Xue M, Tian Y, Sui Y, et al. Protective effect of fucoidan against iron overload and ferroptosis-induced liver injury in rats exposed to alcohol. Biomed Pharmacother 2022;153:113402. [Crossref] [PubMed]
22. Yin J, Hong X, Ma J, et al. Serum Trace Elements in Patients With Polycystic Ovary Syndrome: A Systematic Review and Meta-Analysis. Front Endocrinol (Lausanne) 2020;11:572384. [Crossref] [PubMed]
23. Moon BF, Iyer SK, Hwuang E, et al. Iron imaging in myocardial infarction reperfusion injury. Nat Commun 2020;11:3273. [Crossref] [PubMed]
24. La Rosa P, Petrillo S, Turchi R, et al. The Nrf2 induction prevents ferroptosis in Friedreich's Ataxia. Redox Biology 2021;38:101791. [Crossref] [PubMed]
25. Chen Y, Jiang Z, Li X. New insights into crosstalk between Nrf2 pathway and ferroptosis in lung disease. Cell Death Dis 2024;15:841. [Crossref] [PubMed]
26. Cao J, Chen X, Jiang L, et al. DJ-1 suppresses ferroptosis through preserving the activity of S-adenosyl homocysteine hydrolase. Nat Commun 2020;11:1251. [Crossref] [PubMed]
27. Cao JY, Poddar A, Magtanong L, et al. A Genome-wide Haploid Genetic Screen Identifies Regulators of Glutathione Abundance and Ferroptosis Sensitivity. Cell Rep 2019;26:1544-1556.e8. [Crossref] [PubMed]
28. Hu Z, Mi Y, Qian H, et al. A Potential Mechanism of Temozolomide Resistance in Glioma-Ferroptosis. Front Oncol 2020;10:897. [Crossref] [PubMed]
29. Dodson M, Castro-Portuguez R, Zhang DD. NRF2 plays a critical role in mitigating lipid peroxidation and ferroptosis. Redox Biol 2019;23:101107. [Crossref] [PubMed]
30. Chorley BN, Campbell MR, Wang X, et al. Identification of novel NRF2-regulated genes by ChIP-Seq: influence on retinoid X receptor alpha. Nucleic Acids Res 2012;40:7416-29. [Crossref] [PubMed]
31. Ammal Kaidery N, Ahuja M, Thomas B. Crosstalk between Nrf2 signaling and mitochondrial function in Parkinson’s disease. Mol Cell Neurosci 2019;101:103413. [Crossref] [PubMed]
32. Zhou RP, Chen Y, Wei X, et al. Novel insights into ferroptosis: Implications for age-related diseases. Theranostics 2020;10:11976-97. [Crossref] [PubMed]
33. Zhao J, Lin X, Meng D, et al. Nrf2 Mediates Metabolic Reprogramming in Non-Small Cell Lung Cancer. Front Oncol 2020;10:578315. [Crossref] [PubMed]
34. Sun X, Ou Z, Chen R, et al. Activation of the p62-Keap1-NRF2 pathway protects against ferroptosis in hepatocellular carcinoma cells. Hepatology 2016;63:173-84. [Crossref] [PubMed]
35. Kerins MJ, Vashisht AA, Liang BX, et al. Fumarate Mediates a Chronic Proliferative Signal in Fumarate Hydratase-Inactivated Cancer Cells by Increasing Transcription and Translation of Ferritin Genes. Mol Cell Biol 2017;37:e00079-17. [Crossref] [PubMed]
36. Brown RAM, Richardson KL, Kabir TD, et al. Altered Iron Metabolism and Impact in Cancer Biology, Metastasis, and Immunology. Front Oncol 2020;10:476. [Crossref] [PubMed]
37. Kluknavsky M, Micurova A, Skratek M, et al. A Single Infusion of Polyethylene Glycol-Coated Superparamagnetic Magnetite Nanoparticles Alters Differently the Expressions of Genes Involved in Iron Metabolism in the Liver and Heart of Rats. Pharmaceutics 2023;15:1475. [Crossref] [PubMed]