Adropin as a therapeutic candidate for HFpEF: evidence of oxidative stress mitigation via Nrf2/HO-1 signaling

Abstract

Background

Heart failure with preserved ejection fraction (HFpEF), which accounts for more than half of all heart failure cases worldwide, has emerged as a major public health challenge characterized by substantial morbidity and mortality rates. As adropin is a key regulator of cardiovascular and metabolic homeostasis, this study investigated its therapeutic effects against HFpEF pathogenesis.

Methods

C57BL/6 mice were fed a high-fat diet (60% fat-derived calories) with NG-nitro-L-arginine methyl ester (L-NAME, 0.5 g/L) in drinking water for 8 weeks to induce HFpEF. Adropin-knockout (Ad⁻/⁻) mice were generated, and HFpEF mice received a single intraperitoneal bolus of recombinant adropin (450 nmol kg⁻¹). Cardiac structure and function were quantified by echocardiography. Metabolic status was documented (body weight, fasting glucose, lipids, and systolic/diastolic blood pressure). Myocardial morphology, fibrosis and cardiomyocyte size were examined by hematoxylin‒eosin, Masson’s trichrome and wheat‒germ agglutinin staining. Oxidative stress was evaluated with dihydroethidium fluorescence (ROS) and biochemical assays for malondialdehyde (MDA), superoxide dismutase (SOD) and glutathione (GSH). The protein expression of Nrf2, HO-1, NQO-1 and apoptosis markers (Bcl-2/Bax) was determined by immunoblotting.

Results

HFpEF mice developed significant metabolic disturbances, diastolic dysfunction, myocardial hypertrophy, fibrosis, and increased oxidative stress, alongside markedly reduced serum and myocardial adropin levels. Adropin supplementation improved glucose and lipid metabolism, reduced cardiac hypertrophy and fibrosis, and enhanced diastolic function, whereas adropin knockout exacerbated these pathologies. Mechanistically, adropin activated the Nrf2/HO-1 signaling pathway, increased the expression of antioxidant enzymes (HO-1 and NQO-1), reduced the expression of oxidative stress markers, and regulated apoptosis by increasing Bcl-2 and decreasing Bax expression. HFpEF mice exhibited significant metabolic disturbances, diastolic dysfunction, myocardial hypertrophy, fibrosis, and elevated oxidative stress, alongside markedly reduced serum and myocardial adropin levels. Adropin treatment improved glucose/lipid metabolism, attenuated hypertrophy/fibrosis, and enhanced diastolic function, whereas adropin knockout exacerbated these pathologies. Mechanistically, adropin activated the Nrf2/HO-1 pathway, upregulated antioxidant enzymes (HO-1, NQO-1), reduced oxidative stress markers, and mitigated apoptosis by increasing Bcl-2/Bax ratio.

Conclusion

Adropin improves HFpEF by attenuating metabolic dysregulation, oxidative stress, and myocardial remodeling. Mechanistically, adropin activates the Nrf2/HO-1 pathway, suggesting a novel therapeutic strategy for HFpEF. These findings highlight the potential of adropin to reduce disease burden and improve quality of life in HFpEF patients, addressing critical gaps in current HFpEF management.

Graphical Abstract

Supplementary Information

The online version contains supplementary material available at 10.1186/s12944-025-02703-6.

Introduction

As a prevalent cardiovascular condition, heart failure (HF) persistently accounts for high rates of disease and death worldwide, placing considerable pressure on international health infrastructures [1]. Heart failure with preserved ejection fraction (HFpEF) represents a distinct subtype of HF characterized by a normal or mildly reduced left ventricular ejection fraction (LVEF) accompanied by impaired diastolic function. This dysfunction leads to elevated intracardiac pressures and circulatory abnormalities [1]. The incidence of HFpEF is steadily increasing, particularly among older adults and women. Its pathophysiology is multifactorial and involves alterations in cardiac structure and function, metabolic dysregulation, inflammation, and oxidative stress [2].

Among these contributing factors, oxidative stress plays a pivotal role in the development and progression of HFpEF. In patients with HFpEF, increased oxidative stress is closely associated with cardiomyocyte injury, myocardial fibrosis, and subsequent deterioration of cardiac performance [3]. Consequently, identifying effective strategies to attenuate oxidative stress and preserve cardiac function is critical for improving clinical outcomes in HFpEF patients.

Adropin, a recently identified peptide hormone, has emerged as a potential cardioprotective factor because of its regulatory effects on energy metabolism, inflammation, and oxidative stress [4]. Although it was initially discovered in relation to metabolic homeostasis and insulin sensitivity [5], subsequent studies have revealed its broader physiological significance in cardiovascular health. Notably, circulating adropin levels are reduced in patients with HF and are inversely correlated with HF severity [6]. Moreover, adropin has been shown to ameliorate metabolic and inflammatory disturbances in cardiovascular conditions [7]. A recent study involving 561 type 2 diabetes mellitus patients revealed that individuals with both diabetes and HFpEF exhibited a notable decline in serum adropin levels [8]. However, it remains unclear whether adropin supplementation or activation can confer therapeutic benefits in HFpEF patients. Therefore, further investigation is warranted to elucidate its potential as a treatment modality.

Nuclear factor erythroid 2–related factor 2 (Nrf2), a central transcription factor, plays a pivotal role in modulating antioxidant defense systems by upregulating enzymes such as heme oxygenase-1 (HO-1) [9]. HO-1 exerts antioxidative and anti-inflammatory effects by degrading heme into bioactive molecules with cytoprotective properties, thereby reducing oxidative damage [10]. Activation of the Nrf2/HO-1 signaling pathway has demonstrated promising therapeutic potential in cardiovascular diseases by enhancing the antioxidant defense of cardiomyocytes and mitigating oxidative stress-induced myocardial injury [11]. Recent studies suggest that adropin may activate the Nrf2/HO-1 axis, thereby increasing the intracellular antioxidant capacity and protecting tissues from oxidative damage [12].

Building upon established knowledge of the metabolic regulatory functions of adropin [4–6], this study addresses a critical knowledge gap by investigating its unexplored therapeutic potential in HFpEF. The central hypothesis of this study posits that adropin exerts cardioprotective effects against HFpEF through multimodal mechanisms. The present work provides three fundamental advances in the field: first, causal evidence was established through genetic and pharmacological approaches demonstrating that adropin deficiency exacerbates while supplementation ameliorates HFpEF progression; second, the Nrf2/HO-1 antioxidant pathway, as a novel mechanistic mediator of the cardioprotective effects of adropin, was identified; third, the unique capacity of adropin to simultaneously target multiple pathological axes in HFpEF, a distinct therapeutic advantage over current single-mechanism approaches, was revealed. These findings extend understanding of adropin’s cardiovascular benefits by defining its specific role and mechanisms in HFpEF pathophysiology.

Method

Animals

All animal procedures used male C57BL/6 mice aged 4–6 weeks (Hunan SJA Laboratory Animal Co., China), which were maintained at 22–25 °C under a 12-h light/dark cycle. All animal experiments were conducted under the supervision of the Institutional Animal Care and Use Committee (IACUC) at the Second Affiliated Hospital of Nanchang University, with formal approval from the Medical Research Ethics Committee (approval review no. [2020] A902). Mice were anesthetized with 2% isoflurane inhalation. Euthanasia was performed via cervical dislocation under deep anesthesia, followed by rapid tissue collection.

Ad^-/- mouse generation

CRISPR/Cas9-induced frameshift mutations via nonhomologous end-joining (NHEJ) repair resulted in functional knockout of the Enho gene. Shanghai Model Organisms Center, Inc., successfully generated Ad^-/- mice, which was confirmed by PCR.

HFpEF model construction

In accordance with methods reported in the literature [6, 13], mice in the HFpEF group were fed a HFD with 60% fat content and a solution of 0.5 g/L L-NAME in sterile water for 8 weeks. Establishment of the model was identified on the basis of the diagnostic criteria for HFpEF [14].

Mice fed Adropin protein

In the HFpEF + Ad group, normal mice were fed a high-fat diet and L-NAME in tap water and were intraperitoneally injected with adropin at 450 nmol/kg once a day for 8 weeks. In the HFpEF + Ad^-/- + Ad group, Ad^-/- mice were fed a high-fat diet and L-NAME in tap water and were intraperitoneally injected with adropin at 450 nmol/kg once a day for 8 weeks. The effective concentration of adropin was determined according to a previously described method [5].

Mouse body weight, heart rate and noninvasive blood pressure measurements

The weight of the animals was measured using an electronic scale, and the heart rate and noninvasive blood pressure of the mice were subsequently measured using a small-animal blood pressure meter (Softron, Tokyo, Japan). The mice were placed in a dark container at 37 °C, and their tails were exposed. The pressure sensor was placed on the upper 1/3 of the tail (tail cuff method). When the mouse was stable, the pressure sensor was used to measure and record the blood pressure. The test was repeated three times for each mouse.

Ultrasonic heart examination in mice

After the mice were anesthetized, the following data were collected by ultrasound (Visual Sonics, Toronto, Canada): left ventricular ejection fraction (LVEF); end-diastolic left ventricular diameter (LVID; d); end-diastolic septal wall thickness (IVS; d); left ventricular end-diastolic wall thickness (LVPW; d); left ventricular fractional shortening (LVFS); peak mitral valve flow velocity in the early diastolic period (E); peak mitral valve flow velocity in the late diastolic period (A); mitral valve tissue diastolic velocity in the early diastolic period (e’); and mitral valve tissue diastolic velocity in the late diastolic period (a’). The test was repeated three times for each mouse.

H&E staining and pathological injury scoring of the mouse myocardium

Paraffin-embedded sections of mouse left ventricular tissue were dewaxed, stained with hematoxylin and eosin (H&E), imaged and analyzed. The pathological score of myocardial injury was determined with reference to a scoring method reported in the literature [15]. Injuries included (1) myocardial fiber hypertrophy, degeneration, or necrosis and (2) interstitial hyperemia, edema, inflammatory cell infiltration, or connective tissue hyperplasia. All kinds of lesions received a score of 1, 2, 3 or 4 points according to the degree of severity, ranging from mild to severe, respectively, and a score of 0 points was assigned in the absence of lesions. The sum of the scores was collected. The specific lesions used for the pathological scoring of lung injury were as follows [16]: thickening of the alveolar septum, hemorrhagic infiltration, exudative edema, lymphoid tissue aggregation, and inflammatory cells. Similarly, each lesion was scored on a 0–4 scale, and the sum of the scores was collected.

Masson staining

Myocardial sections from the mice were stained with potassium bichromate, iron hematoxylin, ponceau acid fuchsin, phosphomolybdic acid and aniline blue and then differentiated and dehydrated with glacial acetic acid. Finally, the transparent sections were sealed and photographed. ImageJ was used to assess the extent of the fibrotic changes in the myocardial tissue and measure the areas of collagen deposition.

Wheat germ agglutinin staining

After mouse myocardial tissue sections were dewaxed, they were placed in EDTA solution for antigen repair. Following wheat germ agglutinin (WGA) staining (37 °C, 30 min) and subsequent PBS washes (3×), the sections were counterstained with DAPI (room temperature, 10 min, dark). After mounting, fluorescence images were captured using microscopy.

Detection of blood glucose by ELISA

Glucose quantification was performed in strict accordance with the instructions of the glucose detection kit (Beyotime, Shanghai, China). Briefly, the serum samples were diluted 1:20 with assay buffer to avoid matrix interference. Standard glucose solutions (0–500 mg/dL) and test samples (50 µL each) were assayed in antibody-coated 96-well plates. After the addition of the glucose‒HRP conjugate (50 µL) and incubation (37 °C, 30 min), the plates were subjected to five PBS‒Tween washes. Color development was performed with TMB substrate (100 µL, 15 min, dark), and the reaction was terminated with 2 M H₂SO₄ (50 µL) before optical density measurement at 450 nm. Glucose concentrations were calculated using a 4-parameter logistic standard curve and are expressed as mg/dL.

Detection of blood lipids in serum by ELISA

The experimental methods were carried out according to the instructions of the blood lipid assay kit (Beyotime, Shanghai, China). Briefly, samples were added to an antibody-coated plate and incubated at 25 °C for 2 h. After washing, 100 µL of biotinylated detection antibody was added, followed by 1 h of incubation. Streptavidin-HRP (1:1,000) and TMB substrate were sequentially added, and the absorbance was read at 450 nm.

Detection of serum adropin levels

After frozen mouse serum samples were thawed, the content of adropin was tested according to instructions of the ELISA kit (CUSABIO, Wuhan, China). Serum samples were thawed on ice and diluted (1:2) with buffer before the assay to reduce interference. Standard adropin solutions and test samples were assayed in antibody-coated 96-well plates (100 µL/well, 2 h, 25 °C). After washing (5× PBS-Tween), the plates were incubated with a biotinylated detection antibody (1:1000, 1 h). After incubation with streptavidin-HRP (1:5000, 30 min), color development was performed with TMB (100 µL/well, 15 min, dark) before the reaction was stopped. Measurements were obtained at 450/630 nm (BioTek Synergy H1, Winooski, United States). Adropin concentrations were interpolated from a 4-parameter logistic standard curve and are expressed as pg/mL.

Immunohistochemistry

In accordance with previous reports [17], the immunohistochemistry (IHC) methods performed included tissue dewaxing, antigen repair, the addition of primary and secondary antibodies, chemical staining with DAB and hematoxylin, dehydration and sealing.

Mouse myocardial dihydroethidium staining

The heart was cleaned with cold saline, and sections of the left ventricle were prepared. After rewarming, the sections were incubated with mouse myocardial dihydroethidium (DHE) staining solution in a dark environment for 30 min. After drying, the nuclei were stained with DAPI solution for 10 min. After the sections were cleaned and dried 3 times in a dark environment, an antiquenching agent was added to seal the slices. Finally, the images were observed and photographed with a fluorescence microscope.

Reactive oxygen species detection in mouse cardiomyocytes using the 2,7-dichlorofluorescin diacetate fluorescence probe

After the myocardial tissue was cut into small pieces, enzymatic digestion was carried out, followed by filtration. Then, according to the instructions of the reactive oxygen species (ROS) detection kit, samples were incubated with the 2,7-dichlorofluorescin diacetate (DCFH-DA) fluorescent probe (Beyotime, Shanghai, China). Finally, the fluorescence intensity of the cells in each well of the 96-well plate was determined using an enzyme immunoassay reader at the specified wavelength.

Detection of malondialdehyde by ELISA

The quantification of malondialdehyde (MDA) was performed using a commercial competitive ELISA kit (CUSABIO, Wuhan, China) in strict adherence with the provided instructions. Following tissue homogenization (10% in PBS) and centrifugation (10,000 × g, 15 min, 4 °C), clarified supernatants were diluted 1:10. The ELISA procedure included plate loading with standards (0–100 nM MDA) and samples (50 µL), conjugate/antibody addition (50 µL each), incubation (1 h, 25 °C), washing (4×), substrate reaction (15 min), and termination before absorbance measurement at 450 nm (BioTek Synergy H1, Winooski, United States). The MDA concentrations were interpolated from the standard curve and normalized to the total protein concentration (Bradford assay), and these data are expressed as nM/mg protein.

Measurement of superoxide dismutase activity by ELISA

The superoxide dismutase (SOD) activity was measured by a sandwich ELISA kit (CUSABIO, Wuhan, China) optimized for Cu/Zn-SOD detection. Tissue homogenates were diluted 1:50 with sample buffer. The samples were incubated (2 h, 37 °C), washed, labeled with biotinylated antibody (1 h) and then streptavidin-HRP (30 min). Then, color was developed with TMB (15 min), H₂SO₄ was used to stop the reaction, and the absorbance was read at 450 nm. SOD activity (U/mL) was calculated using a 4-parameter logistic curve. The results were adjusted to total protein and are expressed as U/mg protein.

Quantification of reduced glutathione by ELISA

Following tissue processing (deproteinization with sulfosalicylic acid, centrifugation, neutralization, and dilution), glutathione (GSH) levels were assessed using a competitive ELISA kit (CUSABIO, Wuhan, China). Standard solutions (0–50 µM) and prepared samples (50 µL) were analyzed with conjugate/antibody reagents (50 µL each) in antibody-coated plates (1 h, 25 °C). After washing (5×) and substrate reaction (20 min), the absorbance at 450 nm was measured for quantitative determination, and the results were standardized to tissue parameters.

Immunoblotting

According to previously described methods [17], protein was isolated by electrophoresis. Following wet electrophoretic transfer to PVDF membranes, nonspecific binding sites were blocked by incubation with 5% bovine serum albumin (BSA) for 30 min under ambient conditions. Primary antibodies against Nrf2 (Immunoway, Jiangsu, China), HO-1 (Immunoway, Jiangsu, China), NQO-1 (Cell Signaling Technology, Boston, USA), Bcl2 (Immunoway, Jiangsu, China) and Bax (Immunoway, Jiangsu, China), all diluted 1:1000, were incubated with the samples at 4 °C overnight. After washing, the membranes were probed with diluted fluorescent secondary antibodies (1:10,000) and incubated for 1 h at room temperature in the dark. Images were captured using an Odyssey Clx two-color infrared fluorescence imaging system, and the gray values of the protein bands were analyzed by ImageJ software.

Data analysis

SPSS 19.0 and GraphPad Prism 8.0 were used for statistical analysis. Quantitative data are expressed as the means ± standard deviations. Normality and homoscedasticity were verified prior to statistical analysis. Two-group comparisons were performed by either Student’s t test (equal variances) or the nonparametric Mann‒Whitney U test (unequal variances), whereas multiple-group comparisons were performed by one-way ANOVA. If the variance was homogeneous, the least significant difference (LSD) method was used for analysis; if the variance was uneven, Tamhane’s T2 test was used. A probability value less than 0.05 was regarded as statistically significant.

Results

Serum and myocardial adropin content is decreased in the mouse model of HFpEF induced by a high-fat diet and L-NAME

To establish a murine model of HFpEF, C57BL/6 mice were fed a high-fat diet and administered L-NAME (an NO synthase inhibitor) in their drinking water. The model was validated based on established diagnostic criteria for HFpEF [14]. Compared with control mice, HFpEF mice presented significant increases in body weight, blood glucose, blood lipid levels, and blood pressure (Fig. 1A–D). In addition, HFpEF mice presented an elevated heart rate, increased heart weight, and pronounced myocardial hypertrophy, as indicated by a significantly increased heart weight‒tibial length ratio (Fig. 1E‒G).

Fig. 1.

An HFpEF model was established, and the adropin content was decreased in HFpEF mice. (A) Body weight was increased in HFpEF mice. (B) Blood glucose levels were increased in HFpEF mice. (C) Blood lipid content was increased in HFpEF mice. (D) Blood pressure was increased in HFpEF mice. (E) Heart rate was increased in HFpEF mice. (F) Heart weight was increased in HFpEF mice. (G) The ratio of heart weight to tibial length was increased in HFpEF mice. (H) Diastolic function of the heart was reflected by mitral valve tissue Doppler and blood flow Doppler in early and late mitral valve diastole. (I) Diastolic function was affected, but systolic function of the heart was not affected in HFpEF mice. E/e’: peak systolic velocity of mitral valve flow/diastolic velocity of mitral valve tissue in diastole; E/A: peak systolic velocity of mitral valve flow in diastole/peak diastolic velocity of mitral valve flow in diastole; e’/a’: diastolic velocity of mitral valve tissue in diastole/diastolic velocity of mitral valve tissue in systole; IVS; d: end-diastolic thickness of the interventricular septum; LVID; d: left ventricular end-diastolic diameter; LVPW; d, left ventricular posterior wall end-diastolic thickness; LVFS: left ventricular fractional shortening; LVEF: left ventricular ejection fraction. (J) Cardiac tissue from HFpEF mice was subjected to H&E, Masson, WGA and IHC (adropin antibody) staining. Black arrows: hypertrophic cardiomyocytes; blue arrows: inflammatory cells; red arrows: dead cardiomyocytes; green arrows: denatured cardiomyocytes; brown arrows: fibroblast hyperplasia and myocardial interstitial fibrosis. (K) Myocardial injury scores were analyzed. (L) Myocardial fibrosis was increased in HFpEF mice, as shown by Masson’s trichrome staining. (M) Myocardial hypertrophy was aggravated in HFpEF mice, as shown by WGA staining. (N) Adropin expression was decreased in the HFpEF mouse myocardium, as determined by IHC staining. (O) Serum adropin content was decreased in HFpEF mice. * P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001; ns: not significant

Echocardiographic analysis revealed that HFpEF mice had marked diastolic dysfunction, as evidenced by increased E/e′ and E/A ratios and a decreased e′/a′ ratio. These mice also displayed structural changes consistent with left ventricular hypertrophy, including an increased interventricular septum thickness and posterior wall thickness and a reduced left ventricular internal diameter during diastole. However, systolic function remained unaffected, as indicated by preservation of the LVFS and LVEF (Fig. 1H–I).

Histopathological analyses revealed significant myocardial remodeling in HFpEF mice. H&E staining revealed extensive myocardial damage, characterized by cardiomyocyte hypertrophy, interstitial fibrosis, destruction of the myocardial architecture, inflammatory cell infiltration, and varying degrees of cellular necrosis and degeneration (Fig. 1J–K). Masson’s trichrome staining further confirmed a significant increase in myocardial fibrosis in HFpEF mice (Fig. 1J, L), whereas wheat germ agglutinin (WGA) staining revealed pronounced cardiomyocyte hypertrophy (Fig. 1J, M).

Importantly, IHC analysis revealed a marked reduction in both serum and myocardial adropin levels in the HFpEF model (Fig. 1J, N–O). This finding was corroborated by ELISA, which demonstrated a significant decrease in circulating adropin concentrations in HFpEF mice. Collectively, these results suggest that HFpEF is associated with systemic and myocardial downregulation of adropin, indicating a potential role of adropin deficiency in the pathogenesis of HFpEF.

Adropin knockout does not affect cardiac function or myocardial morphology under normal feeding conditions

To evaluate the baseline effect of adropin deficiency on cardiac structure and function, adropin-knockout (Ad^-/-) mice were used. Immunoblotting confirmed the successful knockout of adropin protein expression in Ad^-/- mouse myocardial tissue, with no detectable expression compared with that in wild-type controls (Fig. 2A–B). To assess whether the absence of adropin affects cardiac function under physiological conditions, a series of cardiac evaluations were conducted in Ad^-/- mice maintained under normal feeding conditions.

Fig. 2.

Cardiac function of Ad^-/- mice under normal feeding conditions. (A, B) Adropin protein expression detected by immunoblotting of Ad^-/- mouse myocardial tissue. **** P < 0.0001. (C) Histopathological examination of heart tissue by H&E staining revealed no significant difference between Ad^-/- mice and normal mice. (D) Cardiac ultrasound indices were not different between normal mice and Ad^-/- mice. (E–F) Myocardial fibrosis in myocardial tissue was not significantly different between Ad^-/- mice and normal mice according to Masson’s trichrome staining; ns: not significant

Histological examination of myocardial tissue using H&E staining revealed no structural abnormalities or morphological differences between Ad^-/- and wild-type mice (Fig. 2C). Cardiac function, assessed by echocardiography, showed no significant changes in key functional parameters, indicating preserved systolic and diastolic function in the absence of adropin (Fig. 2D). Furthermore, Masson’s trichrome staining demonstrated comparable myocardial fibrosis in the two groups, with no evidence of increased collagen deposition in adropin-deficient mice (Fig. 2E–F).

These results indicate that adropin knockout does not adversely affect cardiac morphology or function under baseline, nonpathological conditions. Therefore, the effects of adropin deficiency appear to be context dependent and become apparent primarily under conditions of stress such as HFpEF.

Adropin deficiency aggravates metabolic parameters, blood pressure, and cardiac dysfunction in HFpEF mice

To verify the effects of adropin on HFpEF mice, the following five groups of mice were used: Ad^-/- mice (Ad^-/- mice under normal feeding conditions); HFpEF mice (normal mice fed a high-fat diet and L-NAME in tap water for 8 weeks); HFpEF + Ad^-/- mice (Ad^-/- mice fed a high-fat diet and L-NAME in tap water for 8 weeks); HFpEF + Ad mice (normal mice fed a high-fat diet and L-NAME in tap water and intraperitoneally injected with adropin at 450 nmol/kg once a day for 8 weeks); and HFpEF + Ad^-/-+Ad mice (Ad^-/- mice fed a high-fat diet and L-NAME in tap water and intraperitoneally injected with adropin at 450 nmol/kg once a day for 8 weeks). The results showed that adropin significantly improved body weight in HFpEF mice, whereas adropin knockout further increased the body weight of HFpEF mice, and supplementation with adropin reversed this trend (Fig. 3A). Adropin significantly reduced the blood lipid and blood glucose levels in HFpEF mice, whereas adropin deletion further increased the blood lipid and blood glucose levels in HFpEF mice, and supplementation with adropin reversed these effects (Fig. 3B–C). Adropin improved hypertension in HFpEF mice, whereas adropin knockout further elevated blood pressure in HFpEF mice without affecting heart rate (Fig. 3D–E). Additionally, adropin reduced heart weight in HFpEF mice, whereas heart weight increased in Ad^-/- mice, and treatment with adropin reduced heart weight in HFpEF and Ad^-/-+HFpEF mice (Fig. 3F).

Fig. 3.

Adropin can significantly improve metabolic disorders in HFpEF mice. (A) Adropin affected body weight in HFpEF mice. (B) Adropin affected blood glucose levels in HFpEF mice, as determined by ELISA. (C) Adropin affected the blood lipid content in HFpEF mice, as determined by ELISA. (D) Adropin did not affect the heart rate in HFpEF mice. (E) Adropin affected the heart weight in HFpEF mice. (F) Adropin affected the heart weight in HFpEF mice. (G‒N) Adropin improved the diastolic function of the heart but did not affect systolic function. E/e’: peak systolic velocity of mitral valve flow/diastolic velocity of mitral valve tissue in diastole; E/A: peak systolic velocity of mitral valve flow in diastole/peak diastolic velocity of mitral valve flow in diastole; e’/a’: diastolic velocity of mitral valve tissue in diastole/diastolic velocity of mitral valve tissue in systole; IVS; d: end-diastolic thickness of the interventricular septum, LVID; d: left ventricular end-diastolic diameter; LVPW; d, left ventricular posterior wall end-diastolic thickness; LVFS: left ventricular fractional shortening; LVEF: left ventricular ejection fraction. (O) Adropin improved pathological damage to the myocardium, as shown by H&E, Masson and WGA staining. Black arrows indicate hypertrophic cardiomyocytes; blue arrows indicate inflammatory cells; red arrows indicate necrotic myocardial cells; green arrows indicate degenerated cardiomyocytes; and brown arrows indicate fibroblast hyperplasia and myocardial interstitial fibrosis. (P–R) Pathological myocardial damage, myocardial fibrosis and cardiomyocyte hypertrophy were statistically analyzed. Ad^−/−: Ad^−/− mice under normal feeding condition; HFpEF: normal mice fed a high-fat diet and L-NAME in sterile water; HFpEF + Ad^−/−: Ad^−/− mice fed a high-fat diet and L-NAME in sterile water; HFpEF + Ad: normal mice fed a high-fat diet and L-NAME in sterile water and intraperitoneally injected with adropin at 450 nmol/kg; HFpEF + Ad^−/−+Ad: Ad^−/− mice fed a high-fat diet and L-NAME in sterile water and intraperitoneally injected with adropin at 450 nmol/kg. *P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001; ns: not significant

Echocardiographic evaluation revealed that adropin markedly improved diastolic function, as evidenced by normalization of the E/A, E/e′, and e′/a′ ratios. It also attenuated left ventricular hypertrophy, as indicated by a decrease in the interventricular septal thickness (IVS; d) and posterior wall thickness (LVPW; d) and an increase in the left ventricular internal diameter (LVID; d). In contrast, adropin deficiency worsened diastolic dysfunction and contributed to myocardial remodeling, although it did not significantly alter posterior wall thickness (Fig. 3G–L). Systolic function, as measured by the LVEF and LVFS, remained unchanged across all groups (Fig. 3M–N).

Histological examination using H&E staining confirmed that adropin ameliorated myocardial pathological injury in HFpEF mice. Mice in the HFpEF + Ad group presented reduced cellular necrosis, diminished fibroblast proliferation, and fewer infiltrating inflammatory cells. In contrast, mice in the HFpEF + Ad^-/- group exhibited extensive cardiomyocyte degeneration, interstitial fibrosis, and inflammation. Reintroduction of adropin into adropin-deficient HFpEF mice (HFpEF + Ad^-/- + Ad group) significantly improved myocardial architecture and reduced pathological scores (Fig. 3O–P). Consistently, Masson’s trichrome staining demonstrated that adropin reduced myocardial fibrosis, whereas its absence exacerbated fibrotic remodeling (Fig. 3O, Q). Additionally, WGA staining revealed that adropin attenuated cardiomyocyte hypertrophy, whereas adropin deficiency led to larger cardiomyocyte cross-sectional areas (Fig. 3O, R).

Taken together, these findings indicate that adropin plays a protective role in HFpEF by regulating body weight, glucose and lipid metabolism, blood pressure, myocardial hypertrophy, and fibrosis. Its deficiency exacerbates the pathological manifestations of HFpEF, whereas its supplementation reverses these deleterious effects, underscoring the therapeutic potential of adropin in the treatment of HFpEF.

Adropin alleviates myocardial oxidative stress and regulates the Nrf2/HO-1 signaling pathway in HFpEF mice

Previous studies have shown that ventricular remodeling and diastolic dysfunction in HFpEF are closely associated with elevated levels of cardiac oxidative stress [18]. Myocardial redox homeostasis is maintained through the dynamic interplay of the oxidative and antioxidative systems. ROS and MDA are key indicators of oxidative damage, whereas GSH and SOD are primary components of the antioxidant defense system. An increase in ROS and MDA levels, accompanied by a decrease in GSH and SOD, reflects a pronounced state of oxidative stress.

To assess whether adropin modulates oxidative stress in the myocardium of HFpEF mice, DHE staining and DCFH-DA fluorescence assays were performed to measure myocardial ROS levels in the myocardial tissue of Ad^-/-, HFpEF, HFpEF + Ad^-/-, HFpEF + Ad and HFpEF + Ad^-/-+Ad mice. In addition, the myocardial concentrations of MDA, GSH, and SOD were quantified. The results revealed a significant increase in oxidative stress markers in HFpEF mice. Notably, adropin deficiency further exacerbated oxidative damage, whereas adropin administration attenuated oxidative stress by reducing ROS and MDA levels and restoring GSH and SOD concentrations in myocardial tissue (Fig. 4A–F).

Fig. 4.

Adropin improved myocardial oxidative stress and regulated the Nrf2/HO-1 signaling pathway in HFpEF mice. (A) Adropin reduced the ROS content in fresh-frozen myocardial tissue of HFpEF mice, as shown by DHE staining. (B) ROS levels in fresh-frozen myocardial tissue were statistically analyzed by DHE staining. (C) Adropin reduced the ROS content in fresh cardiomyocytes isolated in vitro, as determined by the DCFH-DA fluorescent probe method. (D–F) Adropin reduced the MDA, SOD and GSH contents in cardiomyocytes, as determined by ELISA. (G) Adropin reduced Nrf2, HO-1, NQO-1, Bcl-2 and Bax-2 protein expression in HFpEF mice, as shown by WB. (H–K) The protein expression of Nrf2, HO-1, NQO-1, Bcl-2 and Bax-2 was quantified. *P < 0.05, ** P < 0.01, ***P < 0.001, ****P < 0.0001

Nrf2 serves as the principal transcriptional regulator of cellular antioxidant responses and is known for its ability to protect tissues from oxidative injury by upregulating the expression of antioxidant enzymes [19]. To determine whether adropin exerts its protective effects through the Nrf2/HO-1 signaling pathway, protein expression in the myocardial tissue of Ad^-/-, HFpEF and HFpEF + Ad^-/- mice was analyzed by immunoblotting. Significant suppression of the Nrf2/HO-1/NQO1 antioxidant pathway was observed in the HFpEF mouse model. This reduction was even more pronounced in HFpEF mice lacking adropin (HFpEF + Ad^-/- group) (Fig. 4G–K). Furthermore, Bcl-2 protein expression was decreased, whereas the expression of the proapoptotic protein Bax was increased, indicating enhanced myocardial apoptosis in the absence of adropin.

Collectively, these findings suggest that adropin mitigates oxidative stress in HFpEF patients by enhancing antioxidant defenses through the Nrf2/HO-1 signaling pathway and by modulating apoptosis-related proteins to reduce myocardial cell death.

Discussion

Adropin’s therapeutic effects were examined in an established HFpEF murine model using two strategies: exogenous adropin administration and endogenous adropin knockout. Consistent with reports of adropin regulating body weight, reducing obesity, and improving lipid profiles [4, 20], adropin treatment improved dyslipidemia (including increased HDL) in HFpEF mice, while its deficiency exacerbated weight gain and dyslipidemia, concomitant with reduced HDL.

Moreover, adropin is an important glucose-regulating factor [21–23]. Here, adropin treatment reduced blood glucose levels in HFpEF mice, while adropin deficiency induced hyperglycemia, supporting its metabolic regulatory role. Previous clinical evidence has revealed that low circulating adropin levels are associated with cardiovascular diseases, including hypertension [24, 25]. In the present study, adropin lowered both systolic and diastolic blood pressure in HFpEF mice without altering heart rate, whereas adropin deficiency increased blood pressure (with unchanged heart rate), indicating adropin’s blood pressure-lowering effect is independent of chronotropic influence.

HFpEF is clinically characterized by preserved left ventricular systolic function with impaired diastolic function and myocardial remodeling [14]. In the model, HFpEF mice exhibited significant left ventricular hypertrophy (increased IVS; d and LVPW; d, reduced LVID; d, P < 0.05), impaired diastolic function (e′/a′ < 1, E/A > 2, decreased E/e′), and preserved systolic function (unchanged LVEF/LVFS) [13], consistent with established HFpEF models and validating the experimental setup. Notably, adropin administration improved diastolic function, attenuated myocardial hypertrophy (via reduced heart weight and improved echocardiographic parameters), and did not affect systolic function. Histopathology further showed adropin reduced cardiomyocyte hypertrophy, myocardial fibrosis, and epicardial fat deposition, highlighting its role in limiting myocardial injury and structural remodeling in HFpEF.

Cardiac oxidative stress is a central contributor to ventricular remodeling and diastolic dysfunction in HFpEF [26]. It reported that adropin deficiency exacerbated the expression of oxidative stress markers [27]. In the model, HFpEF mice presented elevated myocardial ROS and MDA, alongside reduced GSH and SOD, indicating significant oxidative imbalance. Adropin treatment reversed these changes, decreasing ROS/MDA and restoring GSH/SOD levels, demonstrating its crucial role in suppressing myocardial oxidative damage in HFpEF.

Nrf2 is a master regulator of cellular antioxidant responses, primarily through transcriptional activation of antioxidant response element (ARE)-regulated genes, including HO-1 and NAD(P)H: quinone oxidoreductase 1 (NQO1) [28]. Several studies have demonstrated that activation of the Nrf2/HO-1 signaling pathway confers cardioprotective effects by reducing apoptosis, limiting fibrosis, and improving cardiac function [29–32]. For example, Nrf2 activation has been shown to inhibit cardiomyocyte apoptosis via the modulation of Bcl-2 and Bax expression [29]. These findings are in line with previous observations, indicating that the cardioprotective effects of adropin may be mediated, at least in part, through Nrf2/HO-1 activation.

Emerging evidence indicates adropin exerts antioxidative and anti-inflammatory effects via additional pathways, including VEGFR2-mediated ERK1/2 activation [17, 33, 34], suggesting its protective effects involve a complex regulatory network. Mitochondrial dysfunction drives myocardial injury in conditions like sepsis, ischemia-reperfusion, diabetic cardiomyopathy, and heart failure [35, 36], and adropin has been shown to activate Nrf2/ARE signaling, reduce mitochondrial ROS, and enhance antioxidant defenses in septic cardiomyopathy [37]. It also suppresses NLRP3 inflammasome activation and inflammatory cytokines (e.g., TGF-β1 and IL-6) to mitigate immune-mediated myocardial injury [37]. In radiation-induced injury, adropin promotes cardiac recovery via VEGFR2/PI3K/Akt activation, enhancing angiogenesis and reducing apoptosis [38], while also improving eNOS activity, microvascular perfusion, and infarct size [39]. These diverse mechanisms highlight adropin’s therapeutic potential across cardiovascular pathologies involving oxidative and metabolic dysfunction.

Study strengths and limitations

This study has key strengths advancing the understanding of adropin’s therapeutic potential in HFpEF. First, combining genetic knockout and pharmacological supplementation rigorously established adropin’s causal role in HFpEF pathophysiology. Multimodal assessment (echocardiography, histopathology, molecular analyses) strengthened the findings. Importantly, Nrf2/HO-1 pathway was identified as a novel mechanistic link between adropin and oxidative stress mitigation, revealing new therapeutic targets. Adropin’s simultaneous improvement of metabolic parameters and cardiac function underscores its clinical relevance, addressing a critical unmet need. Limitations include the use of a murine model, which may not fully replicate human HFpEF complexity, and the relatively short (8-week) intervention period, limiting insights into chronic progression. While focused on Nrf2/HO-1, other signaling mechanisms likely contribute to adropin’s effects and warrant investigation. Future studies exploring adropin in larger models and human samples, along with optimal dosing/delivery, will enhance translational potential. These limitations highlight research directions without diminishing the current findings’ significance.

Conclusions

Adropin plays a crucial protective role in HFpEF by improving metabolic dysregulation, attenuating myocardial remodeling, and reducing oxidative stress via activation of the Nrf2/HO-1 pathway. These findings position adropin as both a promising therapeutic target and biomarker for HFpEF, particularly in patients with metabolic comorbidities. Future research should translate these findings by evaluating adropin-based therapies in human trials and exploring their synergy with existing HFpEF treatments. This work provides a strong foundation for developing targeted interventions to improve outcomes in HFpEF patients, addressing a critical unmet need.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

Bingda Li, Jingan Rao and Ping Li conceived of the study. Bingda Li, Wansong Hu, Qing Zhou, Yingxing Wu and Fangpeng Liu performed experiments and analyzed date. Bingda Li, Yuxiu Yang and Ping Li wrote the manuscript.

Funding

This work was provided by the Natural Science Foundation of Jiangxi Province (No. 20232BAB206004 to Bingda Li and No. 20242BAB26112 to Ping Li) and the National Natural Science Foundation of China (No. 82460067 to Bingda Li and No. 82460079 to Ping Li).

Data availability

No datasets were generated or analysed during the current study.

Declarations

Consent for publication

All authors are aware of and consent to the publication of the article.

Competing interests

The authors declare no competing interests.