The present retrospective observational study was approved by the Medical Ethics Committee of the First Affiliated Hospital of Hainan Medical University on January 24, 2025 (Approval no. 2025-KYL-016). Sample collection and processing were conducted in strict accordance with the Declaration of Helsinki and the Regulations on the Management of Human Genetic Resources. Written informed consent was obtained from all participants prior to tissue collection.

Calcified aortic valves were collected from patients who underwent aortic valve replacement at the First Affiliated Hospital of Hainan Medical University between January and December, 2024 (CAV group, n=30), and non-calcified aortic valves were obtained from patients with aortic valve regurgitation due to aortic annular dilatation (normal group, n=30). The sex and age characteristics of the patients were as follows: CAV group: 18 males and 12 females; median age, 68 years (range, 56-68 years); Normal group: 17 males and 13 females; median age, 64 years (range, 48-76 years). Following ethics approval on January 24, 2025, archived specimens and corresponding clinical data were retrieved from February 1, 2025, and laboratory analyses commenced on February 17, 2025.

As regards patients in the CAV group, the diagnosis of severe aortic valve calcification (aortic valve area <1.0 cm², transvalvular flow velocity, >4 msec) was confirmed using echocardiography (transthoracic echocardiogram or transesophageal echocardiogram) and cardiac computed tomography, with the exclusion of secondary calcifications, such as those caused by rheumatic heart disease, infective endocarditis, or metastatic calcification associated with malignancies. For the patients in the normal group, the criteria for inclusion were as follows: Echocardiography confirming an aortic valve annulus diameter >27 mm (in males) or >25 mm (in females), absence of calcific lesions on imaging examinations, and preserved leaflet structure and flexibility, as documented in surgical records. Exclusion criteria included bicuspid aortic valve, congenital valve abnormalities, prior valve interventions and advanced chronic kidney disease.

Primary human AVICs (hAVICs) were isolated according to methods described in previous studies (13,14). In brief, the non-calcified aortic valve was collected and washed in sterile phosphate-buffered saline (PBS, Gibco; Thermo Fisher Scientific, Inc.). Subsequently, excess connective tissue and blood vessels were carefully removed using ophthalmic scissors and forceps, leaving only the valve tissue. The processed valve tissue was then cut into small sections of ~1 mm³ and transferred to a new sterile centrifuge tube. Subsequently, 0.25% trypsin was added, and the mixture was incubated at 37°C in a water bath or on a shaker for 15 min. The sample was then centrifuged at 300 × g for 5 min at room temperature (25°C), and the supernatant was discarded. Subsequently, 2 mg/ml collagenase type I solution was added, and the mixture was incubated at 37°C on a shaking incubator for 4 h. Following incubation, an equal volume of DMEM/F-12 medium containing 10% fetal bovine serum (FBS, Gibco; Thermo Fisher Scientific, Inc.) was added. The cell suspension was filtered through a 70-µm cell strainer and centrifuged at 300 × g for 5 min at room temperature (25°C). The supernatant was discarded, and the cells were resuspended in DMEM/F-12 medium (containing 10% FBS). The cell suspension was then seeded into culture dishes and incubated at 37°C with 5% CO₂. When the cells reached 70% confluency, subculture was performed. Cells from passages 3-5 were used in subsequent experiments.

Subsequently, hAVICs at 60-70% confluency were assigned to the following six groups: Control, OM, OM + vector, OM + oe-Sirt1, OM + oe-Sirt1 + ML385, and OM + oe-Sirt1 + Erastin. Specifically, cells in the control group were cultured in standard complete medium containing 10% FBS without osteogenic induction or other interventions. Cells in the OM group were cultured in osteogenic induction medium containing 10 mM β-glycerophosphate, 50 µg/ml ascorbic acid, 100 nM dexamethasone (MilliporeSigma), and 10% FBS until osteogenic differentiation was achieved. Cells in the OM + vector and OM + oe-Sirt1 groups were transfected with the negative control plasmid, pcDNA3.1, and the pcDNA3.1-Sirt1 plasmid (Shanghai GenePharma Co., Ltd.), respectively, using Lipofectamine 3000 (Invitrogen; Thermo Fisher Scientific, Inc.). Following incubation at 37°C for 6-8 h, the medium was replaced with fresh osteogenic induction medium, and the cells were cultured until osteogenic differentiation was complete. Sirt1 overexpression efficiency was verified by RT-qPCR, and the corresponding validation results are provided in Fig. S1A. As for the cells in the OM + oe-Sirt1 + ML385 and OM + oe-Sirt1 + erastin groups, they were transfected with the pcDNA3.1-Sirt1 plasmid as described above; following 6-8 h of incubation at 37°C, the culture medium was replaced with osteogenic induction medium, supplemented with either 1 µM of the Nrf2 inhibitor, ML385 (MedChemExpress; OM + oe-Sirt1 + ML385 group), or 10 µM of the ferroptosis inducer erastin (MedChemExpress, OM + oe-Sirt1 + erastin group); the cells were then cultured until osteogenic differentiation was achieved.

Aortic valve tissues (50 mg), either fresh or stored in liquid nitrogen, were added to 1 ml TRIzol reagent (Invitrogen; Thermo Fisher Scientific, Inc.) and thoroughly homogenized. Total RNA was extracted using the chloroform-isopropanol method as described in the instructions provided with the TRIzol reagent kit, and the extracted RNA was purified with anhydrous ethanol (MilliporeSigma). As for the cell samples, they were washed with PBS and then lysed directly in 1 ml TRIzol reagent. Total RNA was subsequently extracted. The RNA concentration and purity were determined using a NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific, Inc.). Subsequently, 1,000 ng total RNA were reverse transcribed into cDNA using the PrimeScript RT Reagent kit (RR047Q, Takara Bio, Inc.). Equal amounts of cDNA were quantified using TB Green^® Premix Ex Taq™ II FAST quantitative polymerase chain reaction (CN830S, Takara Bio, Inc.). The PCR thermocycling conditions were initial denaturation at 95°C for 30 sec, followed by 40 cycles of denaturation at 95°C for 5 sec and annealing/extension at 60°C for 30 sec. GAPDH was used as the internal reference gene. Data were analyzed using the 2^−ΔΔCq method (15). The primer sequences used are listed in Table I.

Following treatment, the culture supernatants of hAVICs were collected and analyzed for PGC1α (cat. no. KTE61195, Abbkine Scientific Co. Ltd.), mitofusin 2 (MFN2; at. no. KTE61636, Abbkine Scientific Co. Ltd.), optic atrophy 1 (OPA1; at. no. USEE291, Wuhan USCN Business Co., Ltd.), fission 1 (FIS1; at. no. USEJ105, Wuhan USCN Business Co., Ltd.), dynamin-related protein 1 (DRP1; at. no. ab315308, Abcam) levels using commercially available human ELISA kits. All procedures were performed in accordance with the manufacturer's instructions. Briefly, standards and samples were added to pre-coated 96-well plates, followed by incubation at 37°C for 60 min, washing, enzyme conjugation and substrate reaction. The absorbance was measured at 450 nm using a using a Synergy H1 microplate reader (BioTek Instruments, Inc.), and protein concentrations were calculated based on the standard curve. All samples were measured in duplicate, and the average value was used for statistical analysis.

Aortic valve tissues (50 mg) that were previously ground in liquid nitrogen, or treated hAVICs, were lysed with 500 µl radioimmunoprecipitation assay buffer containing 1 mM phenylmethylsulfonyl fluoride (Beyotime Biotechnology). Additionally, cytoplasmic and nuclear proteins were isolated using a nuclear protein extraction kit (cat. no. P0027, Beyotime Biotechnology). The samples were sonicated on ice (10 sec x 3 cycles, with 30-sec intervals). The lysates were centrifuged at 12,000 × g for 15 min at 4°C, and the supernatant was collected. Protein concentrations were measured using the bicinchoninic acid protein assay kit (cat. no. P0012, Beyotime Biotechnolog) according to the standard curve. The protein concentration of all samples was adjusted to 1-2 µg/µl, after which 5X sodium dodecyl sulfate-polyacrylamide gel electropheresis sample buffer (Beyotime Biotechnology) was added, and the samples were denatured in a 95°C metal bath for 5 min. The proteins were separated by 10% sodium dodecyl sulfate-polyacrylamide gel electropheresis (stacking gel: 80 V for 30 min; separating gel: 120 V for 90 min). The proteins were then transferred onto methanol-activated polyvinylidene fluoride (PVDF) membranes by wet transfer at 300 mA for 90 min. The membranes were blocked with 5% skim milk for 1 h at room temperature and subsequently incubated with the corresponding primary antibodies overnight at 4°C. The primary antibodies used were as follows: Sirt1 (1:1,000, cat. no. 9475, Cell Signaling Technology, Inc.), glutathione peroxidase 4 (GPX4; 1:1,000, cat. no. ab125066, Abcam), acyl-CoA synthetase long-chain family member 4 (ACSL4; 1:1,000, cat. no. 9189, Cell Signaling Technology, Inc.), Runt-related transcription factor 2 (RUNX2; 1:1,000, cat. no. ab192256, Abcam), solute carrier family 7 member 11 (SLC7A11; 1:1,000, cat. no. 26864-1-AP, Proteintech Group, Inc.), ferritin heavy chain 1 (FTH1; 1:1,000, cat. no. ab75973, Abcam), Kelch-like ECH-associated protein 1 (KEAP1; 1:1,000, cat. no. 10503-2-AP, Proteintech Group, Inc.), HO-1 (1:1,000, cat. no. 5853, Cell Signaling Technology, Inc.), NAD(P)H quinone dehydrogenase 1 (NQO1; 1:1,000, cat. no. ab80588, Abcam), Nrf2 (1:3,000, cat. no. PA5-27882, Thermo Fisher Scientific, Inc.), β-actin (1:5,000, cat. no. A5441, MilliporeSigma) and histone H3 (1:1,000, cat. no. 4499, Cell Signaling Technology, Inc.). After washing, the membrane was incubated with HRP-conjugated secondary antibodies [anti-rabbit IgG (1:5,000, cat. no. 7074, Cell Signaling Technology, Inc.) and anti-mouse IgG (1:5,000, cat. no. 7076, Cell Signaling Technology, Inc.)] at room temperature for 1 h. Subsequently, enhanced chemiluminescence substrate (cat. no. P0018S, Beyotime Biotechnology) was added to the membrane for 30 sec of incubation. The band grayscale values were quantified using ImageJ software (version 1.53, National Institutes of Health), and the relative expression of the target protein was calculated using β-actin and histone H3 as internal controls.

A total of 1×10⁴ hAVICs were seeded in each well of a 24-well plate. Upon different treatments and osteogenic induction, the cells were washed twice with PBS and then fixed in 4% paraformaldehyde (Gibco; Thermo Fisher Scientific, Inc.) at room temperature for 15 min. Following three additional PBS washes, the cells were subjected to ALP staining and Alizarin Red S staining.

The ALP staining working solution was prepared according to the instructions provided with the ALP Staining kit (C3206, Beyotime Biotechnology). BCIP (5-bromo-4-chloro-3-indolyl phosphate) and NBT (nitro blue tetrazolium), supplied with the kit, were mixed at a 1:1 ratio. Subsequently, 500 µl ALP working solution were added to each well to cover the cells. The cells were then incubated at 37°C for 30 min in the dark until the blue-violet precipitate appeared. Subsequently, the staining solution was discarded, and the cells were washed three times with PBS. Finally, the blue-violet ALP-positive cells were observed and photographed under an optical microscope (Olympus Corporation), and the staining intensity and area were quantified using ImageJ software (version 1.53, National Institutes of Health).

Alizarin Red S staining solution was prepared according to the instructions provided with the Alizarin Red S Staining kit (C0148S, Beyotime Biotechnology). Cells in each well was supplemented with 500 µl Alizarin Red S staining solution and incubated at room temperature for 30 min. The staining solution was then removed, and the cells were washed three times with PBS. The stained cells were then observed and photographed under an optical microscope (Olympus Corporation). Immediately following image acquisition, a 10% citric acid-ammonia buffer (pH 4.1, supplied with the kit) was added to the wells, followed by a 30-min incubation period at room temperature to completely dissolve the staining substance. Lastly, the absorbance at 415 nm was measured using a microplate reader (FLx800, BioTek; Agilent Technologies, Inc.) to quantify the mineralization level.

The cells were seeded in 96- or 6-well plates and treated when they reached a confluency of 60-80%. Following the indicated osteogenic induction and pharmacological/genetic treatments described above, the culture medium was removed, and the cells were gently washed twice with PBS. Subsequently, the total reactive oxygen species (ROS) levels, mitochondrial ROS levels, mitochondrial membrane potential (JC-1), mitochondrial DNA (mtDNA), adenosine triphosphate (ATP) levels and cellular malondialdehyde (MDA), glutathione (GSH), ferrous ion (Fe²⁺) and total iron levels were determined as described below.

Total ROS levels in the cells were measured using 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA, S0034S, Beyotime Biotechnology) and mitochondrial ROS levels were assessed using MitoSOX™ Red (S0061S, Beyotime Biotechnology). The cells were incubated with 10 µM DCFH-DA or 5 µM MitoSOX™ Red at 37°C for 30 min, followed by three washes with PBS. Fluorescence was then detected using a microplate reader (FLx800, BioTek; Agilent Technologies, Inc.) with DCFH-DA [excitation/emission (Ex/Em)=488/525 nm] or MitoSOX™ Red (Ex/Em=510/580 nm).

Relative changes in mtDNA were evaluated by qPCR. Mitochondrial DNA was first extracted using a mitochondrial DNA extraction kit. Mitochondrial COX1 was selected as the target gene, and quantification was performed using SYBR-Green Polymerase Chain Reaction Master Mix (cat. no. A25742; Applied Biosystems; Thermo Fisher Scientific, Inc.). The thermocycling conditions were as follows: Initial denaturation at 95°C for 10 min, followed by 40 cycles of denaturation at 95°C for 15 sec and annealing/extension at 60°C for 1 min. GAPDH was used as the internal control, and data were analyzed using the 2^−ΔΔCq method (15). The primer sequences used were as follows: COX1 forward, 5′-GAA TGC CAC CTT CAT CCG AGA AG-3′ and reverse, 5′-GCT CAC ATT GGA GAA GGA CTC C-3′; and GAPDH forward, 5′-AAG ATC ATC AGC AAT GCC TCC-3′ and reverse, 5′-AGG TTT TTC TAG ACG GCA GG-3′.

The ATP content in the cells was measured using the ATP assay kit (cat. no. S0026, Beyotime Biotechnology). Specifically, ATP lysis buffer was added to a 6-well plate and the cells were lysed on ice for 5 min. The lysate was then centrifuged at 10,000 g for 10 min at 4°C. Immediately following centrifugation, 50 µl of the supernatant were mixed with 50 µl of the fluorescent substrate and incubated at room temperature for 5 min. The ATP levels were then measured using a microplate reader (FLx800, BioTek; Agilent Technologies, Inc.) at Ex/Em=485/520 nm, and the ATP concentration was calculated based on the standard curve.

The levels of MDA, GSH, Fe²⁺ and total iron were measured according to the instructions provided with the MDA assay kit (cat. no. S0131M, Beyotime Biotechnology), GSH assay kit (cat. no. S0053, Beyotime Biotechnology), Fe²⁺ assay kit (cat. no. BC5415, Beijing Solarbio Science & Technology Co., Ltd.) and total iron assay kit (cat. no. BC5315, Beijing Solarbio Science & Technology Co., Ltd.). Briefly, cells from a 6-well plate were harvested and lysed, after which reaction solutions were added. The absorbance was then measured using a microplate reader (FLx800, BioTek; Agilent Technologies, Inc.), and the MDA, GSH, Fe²⁺ and total iron levels were calculated based on the standard curve.

Mitochondrial membrane potential was measured using the JC-1 mitochondrial membrane potential assay kit (cat. no. C2003S, Beyotime Biotechnology). The cells were incubated with JC-1 working solution at 37°C for 20-30 min, followed by three washes with PBS. The ratio of red fluorescence (Ex/Em=550/600 nm) to green fluorescence (Ex/Em=488/525 nm) was detected using a microplate reader (FLx800, BioTek; Agilent Technologies, Inc.).

All animal experiments in the present study were conducted in accordance with the guidelines outlined in the Guide for the Care and Use of Laboratory Animals (National Institutes of Health Publication No. 85-23, 2011 revision) (16) and approved by the Ethics Committee of Hainan Medical University (Approval no. HYALL-2025-028). Male apolipoprotein E-deficient (Apoe^−/−) mice (8 weeks old, weighing 20-25 g; n=30) were purchased from the Guangdong Provincial Laboratory Animal Center. Mice were housed under specific pathogen-free conditions at 22±2°C and 50-60% relative humidity, with a 12/12-h light/dark cycle and ad libitum access to food and water.

Following acclimatization, mice were randomly assigned to the following three groups (n=10 per group): Normal, calcified and calcified + SRT2104. The mice in the normal group received standard laboratory chow (Beijing Keao Xieli Feed Co., Ltd. China), consisting primarily of corn, soybean meal, fish meal, wheat flour, yeast powder, vegetable oil, vitamins and minerals, with ≥18% crude protein and ≥4% crude fat. The mice in the calcified group received a Western diet (Beijing Keao Xieli Feed Co., Ltd.) containing 21% fat and 0.15% cholesterol. The mice in the calcified + SRT2104 group received the same Western diet supplemented with SRT2104, a Sirt1 agonist (MedChemExpress), at 1.33 g/kg diet, corresponding to ~100 mg/kg/day based on the average daily food intake per mouse under standard housing conditions.

For mechanistic validation, recombinant AAV9 vectors carrying shRNAs targeting mouse HO-1 or FTH1, together with the corresponding negative-control vector (AAV-NC), were commercially constructed, packaged, purified and titrated by GeneChem Co., Ltd. The shRNA sequences used for HO-1 and FTH1 silencing are provided in Table SI. The purified viral preparations were supplied at a titer of ~1×10¹² vg/ml. Each mouse received 1×10¹¹ vg in a final volume of 100 µl via tail-vein injection. Successful knockdown was subsequently verified in mouse aortic valve tissues by measuring HO-1 and FTH1 mRNA expression (Fig. S1).

All mice were housed in a Specific Pathogen-Free (SPF) facility under a 12/12-h light/dark cycle. The experimental period lasted for 24 weeks. For terminal tissue collection, anesthesia was induced with 3-4% isoflurane and maintained with 1.5-2.0% isoflurane throughout the procedure. At the experimental endpoint, mice were euthanized by inhalation of 5% isoflurane until complete loss of the pedal withdrawal reflex and spontaneous respiration. Death was confirmed by the absence of both heartbeat and respiration before tissue collection, in accordance with institutional animal care and ethical guidelines.

After completing the experimental protocol, mice were anesthetized with 2.5% isoflurane, followed by left ventricular perfusion with PBS and 4% paraformaldehyde for tissue fixation. The heart and aorta were then excised and further fixed in 4% paraformaldehyde at 4°C for 24 h. Following washing with PBS, the tissues were immersed in 15% sucrose at 4°C for 12 h, and then transferred to 30% sucrose at 4°C for a further 12 h for gradient dehydration. Subsequently, the tissues were embedded in optimal cutting temperature compounds, frozen at -20°C, and sectioned at a thickness of 5 µm. For H&E staining, the frozen sections were brought to room temperature, washed with PBS, and stained with hematoxylin and eosin staining solutions (Wuhan Servicebio Technology Co., Ltd.). Sections were stained with hematoxylin for 5 min at room temperature, rinsed with running water, differentiated and blued, and subsequently counterstained with eosin for 2 min at room temperature. After washing, the sections were dehydrated, cleared and mounted. Histopathological changes in the aortic valve leaflets were examined and photographed using a light microscope (Olympus Corporation).

Statistical analyses were performed using SPSS version 26.0 (IBM Corp.) and graphs were generated with GraphPad Prism 9.2.0 (GraphPad Software). Data distribution was assessed using the Shapiro-Wilk test for normality and Levene's test for homogeneity of variance. All data are presented as the mean ± standard deviation (SD). For comparisons between two groups, an unpaired two-tailed Student's t-test was applied. For comparisons among multiple groups, one-way analysis of variance (ANOVA) followed by Tukey's post hoc test was used. A value of P<0.05 was considered to indicate a statistically significant difference.

To explore the role of Sirt1 in CAVD, the present study first examined its expression in human aortic valve tissues and in vitro osteogenic models. The results of RT-qPCR and western blot analysis disclosed that, compared with the normal group, Sirt1 expression was significantly lower in the aortic valve tissue of the CAV group (Fig. 1A and B). Similarly, osteogenic stimulation significantly reduced the expression of Sirt1 in cultured hAVICs relative to the control group (Fig. 1C and D). These data suggested the consistent downregulation of Sirt1 in both clinical and experimental calcification models.

Given the reduced expression of Sirt1 in calcified settings, the present study then investigated whether restoring its expression could influence osteogenic transition. To further investigate the association between Sirt1 and CAVD, Sirt1 expression was increased in hAVICs in vitro using an overexpression vector (pcDNA3.1-Sirt1) (Fig. 2A). The cells were subjected to osteogenic induction, and ALP and Alizarin Red S staining were carried out when the cells reached full maturation. The experimental results revealed that in comparison with the control group, the cells in the OM group exhibited a significantly higher ALP activity, enhanced mineralization and the increased formation of calcified nodules. By contrast, the OM + oe-Sirt1 group displayed a significantly reduced ALP activity, lower mineralization and decreased calcified nodule formation compared with the OM + vector group (Fig. 2B and C). Consistently, the calcifying conditions markedly increased nuclear RUNX2 accumulation, whereas Sirt1 overexpression significantly reduced nuclear RUNX2 levels, indicating the inhibition of RUNX2 nuclear translocation (Fig. 2D). RT-qPCR revealed elevated mRNA levels of the osteogenic markers, RUNX2, OPN, osterix and ALP in the OM group, which were all significantly downregulated in the OM + oe-Sirt1 group (Fig. 2E). These data demonstrated that the overexpression of Sirt1 inhibited the calcification of osteogenically-induced hAVICs.

Recent research has proposed that alterations in mitochondrial metabolism are a key determinant in the development of CAVD (17). Based on this, herein, it was hypothesized that Sirt1 may serve as a critical link between CAVD and mitochondrial function. As shown in Fig. 3A and B, the overexpression of Sirt1 significantly decreased both total and mitochondrial ROS levels in hAVICs compared to the OM + vector group. Correspondingly, JC-1 assay revealed that mitochondrial membrane potential, which was reduced in the OM-treated cells, was markedly restored by the overexpression of Sirt1 (Fig. 3C). In line with this, the mtDNA content and intracellular ATP levels were also significantly elevated in the OM + oe-Sirt1 group (Fig. 3D and E). ELISA revealed that the overexpression of Sirt1 reversed the OM-induced downregulation of PGC1α, MFN2 and OPA1, and suppressed the upregulation of FIS1 and DRP1, indicating improved mitochondrial dynamics (Fig. 3F-J). These results suggested that the overexpression of Sirt1 improved mitochondrial function under conditions of osteogenic stress.

Additionally, it has previously been demonstrated that ferroptosis plays a critical role in CADV (5). Notably, Sirt1, mitochondria and ferroptosis are often interrelated (18). Therefore, in the present study, it was hypothesized that Sirt1 may induce ferroptosis in cells. To validate this hypothesis, the levels of MDA, GSH, Fe²⁺ and total iron in hAVICs were measured following the overexpression of Sirt1. The results revealed that the overexpression of Sirt1 significantly reduced the MDA, Fe²⁺ and total iron levels, while it significantly restored the GSH levels in the OM-treated hAVICs (Fig. 4A-D). In parallel, the results of western blot analysis revealed that the overexpression of Sirt1 increased the expression of the anti-ferroptotic proteins, GPX4, SLC7A11 and FTH1, and suppressed the pro-ferroptotic marker, ACSL4, compared to the OM + vector group (Fig. 4E and F). These findings suggest that Sirt1 inhibits ferroptosis in hAVICs under conditions of osteogenic stress.

Previous studies have demonstrated that Sirt1 activates the Nrf2/HO-1 pathway through its deacetylase activity, thereby inhibiting ferroptosis (19,20). Therefore, it was hypothesized that the Nrf2/HO-1 pathway may be the key signaling pathway responsible for the induction of ferroptosis following the reduced expression of Sirt1 in CAVD. Western blot analysis revealed that, relative to the control group, the OM group exhibited a significant upregulation in KEAP1 protein expression, a marked downregulation in the levels of HO-1 and NQO1 proteins, and a significant increase in the Nrf2 protein levels in the cytoplasm, while the nuclear Nrf2 protein levels were significantly decreased. Conversely, compared to the OM + vector group, the OM + oe-Sirt1 group had significantly lower KEAP1 levels, higher HO-1 and NQO1 levels, as well as reduced Nrf2 levels in the cytoplasm, and elevated Nrf2 levels in the nucleus (Fig. 5). These findings suggest that the overexpression of Sirt1 activates the Nrf2/HO-1 pathway in osteogenically-induced hAVICs.

To verify whether the anti-calcification effects of Sirt1 are dependent on Nrf2/HO-1 activation and the suppression of ferroptosis, hAVICs overexpressing Sirt1 were treated with Nrf2 inhibitor (ML385) or ferroptosis inducer (erastin). Initially, ALP staining and Alizarin Red S staining revealed that, compared to the OM + oe-Sirt1 group, the OM + oe-Sirt1 + ML385 and OM + oe-Sirt1 + erastin groups exhibited a significantly increased ALP activity, mineralization and calcified nodule formation (Fig. 6A and B). Consistently, the expression levels of the osteogenic markers, RUNX2, OPN, osterix and ALP, were upregulated (Fig. 7A). Moreover, the levels of the mitochondrial function markers, PGC1α, MFN2 and OPA1, were downregulated, whereas the levels of FIS1 and DRP1 were upregulated (Fig. 7B). Additionally, in comparison with the OM + oe-Sirt1 group, the OM + oe-Sirt1 + ML385 and OM + oe-Sirt1 + erastin groups exhibited significantly reduced levels of GPX4, SLC7A11 and FTH1, and a significant increase in ACSL4 protein expression (Fig. 7C). These data indicated that the inhibition of the Nrf2/HO-1 pathway or the promotion of ferroptosis enhanced the calcification of osteogenically-induced hAVICs.

To further investigate the roles of HO-1 and FTH1 in the Sirt1-mediated anti-calcification effects, hAVICs overexpressing Sirt1 were subjected to HO-1 or FTH1 knockdown under osteogenic conditions.

Alizarin Red S and ALP staining demonstrated that the overexpression of Sirt1 markedly reduced mineralization and osteogenic activity, whereas silencing either HO-1 or FTH1 partially reversed these inhibitory effects (Fig. 8A and B). Both interventions increased calcified nodule formation and ALP activity compared with the Sirt1 overexpression group.

At the transcriptional level, the overexpression of Sirt1 significantly increased Sirt1 and HO-1 mRNA expression, whereas HO-1 knockdown effectively reduced the HO-1 levels. By contrast, FTH1 expression was decreased following FTH1 silencing, confirming the efficiency of gene-specific knockdown (Fig. 8C). Consistently, the expression of osteogenic markers, including RUNX2, OPN, osterix and ALP, was significantly suppressed by Sirt1 overexpression, but was restored upon the knockdown of either HO-1 or FTH1 (Fig. 8D).

These findings were further supported by western blot analysis, which revealed an increased nuclear RUNX2 accumulation following HO-1 or FTH1 silencing compared with the Sirt1 overexpression group (Fig. S2A). To further investigate the underlying signaling pathway, key components of the Nrf2/HO-1 axis were analyzed. Compared with the OM and OM + vector + shNC groups, Sirt1 overexpression markedly reduced KEAP1 protein expression and increased the nuclear Nrf2 levels, accompanied by a decrease in cytoplasmic Nrf2, and the upregulation of HO-1 and NQO1 expression. Upon HO-1 knockdown, HO-1 protein expression was significantly reduced, and NQO1 levels were correspondingly decreased. In addition, KEAP1 expression and both nuclear and cytoplasmic Nrf2 levels exhibited no obvious changes compared with the Sirt1 overexpression group. By contrast, FTH1 silencing did not significantly affect the expression of HO-1, NQO1, KEAP1, or the nuclear and cytoplasmic distribution of Nrf2 compared with the Sirt1 overexpression group (Fig. S2B).

In addition, biochemical analysis revealed that Sirt1 overexpression reduced MDA levels and increased the GSH content, while these effects were reversed by the knockdown of either HO-1 or FTH1 (Fig. S2C), indicating a restoration of oxidative stress and ferroptosis-related imbalance. Collectively, these results indicate that both HO-1 and FTH1 are involved in mediating the inhibitory effects of Sirt1 on osteogenic differentiation and ferroptosis in hAVICs.

To validate the in vitro findings in vivo, a mouse model of CAVD was established using Apoe^−/− mice fed a Western diet, with or without the administration of SRT2104, a pharmacological Sirt1 activator. Following 24 weeks of feeding, the tissue sample was collected, and H&E staining was performed for histological evaluation.

The histological evaluation of aortic valve tissues via H&E staining revealed that the calcified tissues exhibited marked valve thickening, structural disorganization and inflammatory cell infiltration. By contrast, the SRT2104-treated group demonstrated preserved trilaminar architecture, reduced leaflet thickness and diminished inflammatory features, suggesting alleviated pathological remodeling (Fig. 9A). These findings indicate that SRT2104 ameliorates calcification-associated structural changes in vivo.

At the molecular level, RT-qPCR analysis revealed that the levels of osteogenic markers (RUNX2, OPN, osterix and ALP) were significantly upregulated in the calcified group and were markedly reduced following SRT2104 treatment (Fig. 9B), indicating the suppression of osteogenic transformation.

To determine whether the protective effects were mediated via the Nrf2/HO-1 pathway, the levels of related proteins were assessed. Compared to the calcified group, the SRT2104-treated mice exhibited reduced KEAP1 levels, an elevated expression of HO-1 and NQO1, and an enhanced nuclear accumulation of Nrf2, along with decreased cytoplasmic retention (Fig. 9C and D). Collectively, these findings suggest that the pharmacological activation of Sirt1 attenuates calcification-associated pathological remodeling in vivo, at least in part, through the reactivation of Nrf2/HO-1 signaling and the suppression of osteogenic transformation.

To validate the relative contributions of HO-1 and FTH1 in vivo, Apoe^−/− mice treated with SRT2104 were further subjected to AAV-mediated silencing of HO-1 or FTH1. Histological analyses, including H&E and Alizarin Red S staining, revealed that SRT2104 markedly attenuated aortic valve pathological remodeling and calcium deposition compared with the calcified group. The silencing of either HO-1 or FTH1 partially reversed these protective effects, as evidenced by increased leaflet thickening, structural disorganization, inflammatory infiltration and Alizarin Red-positive staining. The AAV-shHO-1 and AAV-shFTH1 groups exhibited comparable pathological changes and calcium deposition (Fig. 10A and B).

At the molecular level, RT-qPCR demonstrated that SRT2104 significantly reduced the expression of osteogenic markers (RUNX2, OPN, osterix and ALP), whereas the knockdown of HO-1 or FTH1 partially restored their expression levels (Fig. 10C). Consistently, western blot analysis revealed that SRT2104 decreased nuclear RUNX2 protein expression, while this effect was reversed following the silencing of HO-1 or FTH1 (Fig. S3A).

Further analysis of the Nrf2/HO-1 signaling pathway revealed that SRT2104 treatment decreased KEAP1 expression, increased nuclear Nrf2 levels, reduced cytoplasmic Nrf2 levels, and upregulated the expression of HO-1, NQO1 and FTH1. Notably, FTH1 expression was reduced in calcified mice compared with the normal controls and was restored following SRT2104 treatment. Upon HO-1 knockdown, HO-1 protein expression was markedly reduced, accompanied by decreases in the NQO1 and FTH1 levels, whereas KEAP1 expression and Nrf2 nuclear and cytoplasmic distribution exhibited no obvious changes compared with the SRT2104-treated group. By contrast, FTH1 silencing markedly reduced FTH1 protein expression, while the KEAP1, Nrf2 (nuclear and cytoplasmic), HO-1 and NQO1 levels remained largely unaltered compared with the SRT2104-treated group (Fig. S3B). Collectively, these results indicate that both HO-1 and FTH1 participate in the protective effects of Sirt1 activation against aortic valve calcification in vivo.