Mitochondrial Iron-Induced Cell Death in Blood Vessel Muscle Linked to Atherosclerosis and Influenced by NCOA4 and Gualou-Xiebai

ox-LDL (YB001), Anti-NCOA4, Anti-GPX4, were obtained from Immunoway (YT0302, YN3047, Beijing, China). Anti-SLC7A11 (26864-1AP), Anti-GAPDH (TA-08). Gualou and Xiebai were purchased from Beijing Tong Ren Tang Co., Ltd. (Beijing, China; batch number: 211001, 211001). An ELISA kit was obtained from Jianglai Biotechnology (Shanghai, China). Erastin was purchased from SparkJade (SJ-MX0039, Jinan, China).

Ninety male ApoE^−/− mice and ten male wild-type C57BL/6J mice (6–8 weeks old, 20–22 g) were obtained from Jiangsu Hua Chuang Xin Nuo Pharmaceutical Technology Co., Ltd. (Taizhou, China). Following a one-week acclimatization period, the ApoE^−/− mice were randomly assigned to five experimental groups (n = 10 per group), while the C57BL/6J mice served as the genetic control group. Mice were maintained under specific pathogen-free (SPF) conditions in a controlled environment (temperature 22 ± 2 °C, humidity 50 ± 10%, 12 h light/12 h dark cycle). They were housed in standard ventilated cages (maximum 5 mice per cage) with corn cob bedding and provided ad libitum access to the respective experimental diets (normal chow or high-fat diet) and autoclaved drinking water. All procedures for animal experiments were approved by the Ethical Review Committee for Laboratory Animal Welfare of Anhui University of Chinese Medicine.

C57BL/6 J mice (n = 10) were fed with a normal diet, and ApoE^−/− mice were fed with a high-fat diet (40% kcal fat, 20% kcal protein, 40% kcal carbohydrate, Cat # D12108C) to simulate an AS model. All animal experiments were approved by the Ethics Committee of Anhui University of Chinese Medicine (approval number: AHUCM-mouse-2023011, approval date: 15 October 2023). The animal experiments were conducted in accordance with the ethical standards established by the Laboratory Animal Welfare Committee of Anhui University of Traditional Chinese Medicine.

Mice were randomly assigned to groups and administered tail vein injections of the vehicle control (PBS with 0.1% BSA) at week 4 and week 8, and the observation period continued until week 16. At 16 weeks of age, mice were fasted for 12 h prior to euthanasia to minimize metabolic variability. Euthanasia was induced via intraperitoneal injection of sodium pentobarbital (50 mg/kg BW), a method approved by the Ethical Review Committee for Laboratory Animal Welfare of Anhui University of Chinese Medicine. Anesthesia depth was monitored by assessing the loss of corneal reflex and response to a painful toe pinch. Upon confirmation of absent vital signs for over 5 min, cervical dislocation was performed as a secondary step to ensure euthanasia. Immediately after confirmation of death, blood was collected via retro-orbital puncture, and tissues were harvested for subsequent analyses. Serum aliquots were stored at −80 °C without repeated freeze–thaw cycles following centrifugation; aortas were snap-frozen in liquid nitrogen and stored at −80 °C for molecular analyses or fixed in 4% paraformaldehyde for histological examinations. These procedures have been supplemented in the relevant Methods section to ensure reproducibility. The ApoE^−/− mice were randomly assigned to the following treatment groups (n = 10): (a) high-fat diet (HFD); (b) HFD + LvNCOA4; (c) HFD + LvNC; (d) HFD + shNCOA4; (e) HFD + shNC; GLXB-L (3 mg/kg) group, GLXB-M (6 mg/kg) group, GLXB-H (12 mg/kg) group, and simvastatin (5 mg/kg) group. The drugs were given once daily by oral gavage for 4 weeks. Finally, serum and aorta were collected for further analysis.

To achieve NCOA4 overexpression and knockdown in atherosclerotic (AS) mice, we utilized lentiviral vectors from Gemma Biologicals (Shanghai, China), specifically Lv18E-NCOA4 (for overexpression) and NCOA4-MUS-1030/149/1345 (for knockdown), along with their respective controls. Lv18E-NCOA4 was constructed by cloning full-length mouse NCOA4 cDNA (NCBI GenBank accession no. NM_001145263.2↗) into the pCDH-CMV-MCS-EF1-copGFP lentiviral backbone, with the empty vector control (LvNC) using the same backbone without the NCOA4 insert; the genomic structure of the overexpression vector was CMV-MCS-hPGK-GFP-T2A-Puro. For knockdown, shNCOA4 and non-targeting scrambled control (shNC) were generated using the pLKO.1-puro lentiviral shRNA vector (Gemma Biologicals, catalog no. SHC001), with the shNCOA4 target sequence 5′-GCAACGUGAAGACCUACAATT-3′ and shNC showing no predicted homology to mouse transcripts. ApoE^−/− mice were administered 80 μL of each virus (titer: 1 × 10⁹ TU/mL) or their corresponding empty vectors via tail vein injection [22].

The GLXB herbal pair, composed of Gualou (Trichosanthes kirilowii Maxim) and Xiebai (Allium macrostemon Bunge), was procured from Beijing Tongrentang Co., Ltd. (Batch numbers: 211001, 211001, Beijing, China). The crude herbs at a weight ratio of 2:1 (Gualou:Xiebai; 100 g:50 g) were pulverized and subsequently macerated in five volumes of 50% aqueous ethanol for 1 h. Extraction was performed twice via reflux extraction, with each session lasting 2 h, using five volumes of 50% ethanol. The resulting extracts were pooled, concentrated to a final concentration of 0.6 g crude herb per milliliter, and stored at 4 °C [24]. All GLXB extracts were administered via oral gavage (intragastric administration) at a regimen of once daily, with consecutive administration for a total duration of 4 weeks.

After one week of acclimatization, 20 Sprague Dawley rats were randomly assigned to a normal serum group (n = 10) and a GLXB serum group (n = 10). Rats in the GLXB group were orally administered the concentrated extract at a dose of 20 g/kg. One hour after the last administration, blood was collected from the abdominal aorta. The samples were incubated at 37 °C for 30 min and then centrifuged at 3000 rpm and 4 °C for 15 min. The supernatant was collected, filtered through a 0.22 μm membrane, and stored at −80 °C for subsequent experiments. The in vitro administration regimen was set as a co-incubation with 50 μg/mL ox-LDL and GLXB serum at a concentration of 24 h [22].

Mouse vascular smooth muscle cells were maintained in high-glucose DMEM supplemented with 10% FBS and 1% penicillin–streptomycin at 37 °C with 5% CO₂. After serum starvation, VSMCs were divided into 12 groups. The control group was cultured in 10% DMEM, while the ox-LDL group was stimulated with 50 μg/mL ox-LDL for 24 h. The ox-LDL + Fer-1 group received 4 h pre-treatment with 1 μmol/L Fer-1 followed by 24 h co-incubation with ox-LDL. The ox-LDL + Erastin group was co-treated with 10 μmol/L Erastin and ox-LDL for 24 h. siRNA transfection groups (siNC, siNCOA4, siFTMT, and co-transfected siNCOA4 + siFTMT) were stimulated with ox-LDL 48 h post-transfection for 24 h. The ox-LDL + siNCOA4 + Erastin group was co-treated with Erastin and ox-LDL post-transfection. The ox-LDL + MitoQ group was pre-treated with 100 nmol/L MitoQ for 2 h before ox-LDL co-incubation. The MitoVES-containing groups (ox-LDL + MitoVES and ox-LDL + siNCOA4 + MitoVES) were treated with 5 μmol/L MitoVES and ox-LDL for 24 h. ox-LDL + GLXB-L (5%)/GLXB-M (10%)/GLXB-H (15%) groups (VSMCs stimulated with 50 μg/mL ox-LDL and cultured with 5, 10, and 15% GLXB-containing serum), ox-LDL + Lv-LOX-1 group (LOX-1 receptor overexpress VSMCs cultured with 50 μg/mL ox-LDL), ox-LDL+Lv-LOX-1 + GLXB-H group (LOX-1 receptor overexpress VSMCs cultured 15% GLXB-containing serum containing 50 μg/mL ox-LDL for 24 h [27].

After overnight incubation of VSMCs in a 6-well plate, the control plasmid was transfected using Lipofectamine 2000 (Solarbio, Beijing, China), 6 h post-transfection, the culture medium was replaced, and the cells were treated with ox-LDL for 24 h. Subsequently, VSMCs were transfected with either siNCOA4 constructs (siNCOA4-1, siNCOA4-2, siNCOA4-3) or an NCOA4 overexpression lentivirus (Lv18E; CMV-MCS-hPGK-GFP-T2A-Puro). A concentration of 5 μg/mL Polybrene was employed for the transfection of the Lv18E lentivirus [22].

The mouse eyeball was dissected with scissors along the midline of the abdomen, and the aorta was bluntly separated and removed in normal saline. Saline was injected into the heart to flush the blood vessels to remove excess blood. Subsequently, the adipose tissue around the aorta was carefully removed using microscissors and fibrous tweezers. Finally, the lipid accumulation in the mouse aorta was detected and photographed by stereomicroscope (SteREO Discovery V20, Zeiss, Jena, Germany).

Aortic tissues were fixed, paraffin-embedded, and sectioned at 5 μm. Sections were deparaffinized in xylene and rehydrated through a graded ethanol series (100%, 95%, 80%, and 70%; 2 min each). After washing, staining was performed with hematoxylin (3–8 min), differentiated in 1% acid ethanol, blued in 0.6% ammonia, and counterstained with eosin (1–3 min). Sections were then dehydrated through a reverse ethanol series (70%, 80%, 95%, and 100%; 2 min each), cleared in xylene, and mounted for microscopic (BX53, Olympus, Tokyo, Japan) examination and ImageJ analysis.

After washing with PBS, the tissue sections were blocked for 2 h in PBS containing 0.3% Triton X-100 and 0.5% bovine serum albumin. The primary antibodies were prepared and incubated at 4 °C overnight. The secondary antibody was incubated for 2 h. DAPI was dropped and incubated in the dark for 15 min. PBS was added, and the washing was repeated 3 times. They were observed and photographed using a laser confocal microscope.

Well-grown VSMCs were seeded in 6-well plates, and the reaction was continued for 1.5 h in the cell incubator after the addition of EdU working solution. After washing with PBS buffer, 1 mL of 4% PFA was added and fixed for 15 min at room temperature. An additional 1 mL of permeabilization solution (0.3% Triton X-100 in PBS) was added and incubated for 15 min at room temperature. A click reaction solution was prepared and reacted at room temperature in the dark for 30 min. DAPI was dropped and incubated in the dark for 15 min and washed with PBS. They were observed and photographed using a fluorescence microscope.

Aortic vascular smooth muscle cells (VSMCs) were seeded at 1 × 10⁶ cells per well in 6-well plates and cultured until 100% confluence. A sterile 200 μL pipette tip was used to create uniform vertical scratches across the cell monolayer, and detached cells were removed by washing three times with PBS. Subsequently, cells were subjected to experimental treatments (consistent with in vitro treatment conditions described in Section 2.5). Images of the scratch areas were captured at 0 h and 24 h post-injury using an inverted phase-contrast microscope (Olympus Inc., Japan). The scratch healing rate was calculated as [(scratch width at 0 h − scratch width at 24 h)/scratch width at 0 h] × 100% using ImageJ software 1.52v.

VSMCs were seeded at 1 × 10⁴ cells per well in the upper chamber of 24-well transwell inserts (8 μm pore size, Labgic, Beijing, China) containing serum-free DMEM. The lower chamber was filled with 600 μL complete DMEM (10% fetal bovine serum). The same three experimental treatments as above were applied to the upper chamber medium. After 24 h of incubation at 37 °C with 5% CO₂, non-migrated cells on the upper surface of the membrane were gently removed with a cotton swab. Migrated cells adhering to the lower surface were fixed with 4% paraformaldehyde for 15 min, stained with 0.5% crystal violet solution for 10 min, and rinsed three times with PBS to remove excess dye. Images of migrated cells were acquired from five random fields per insert using an inverted phase-contrast microscope (Olympus Inc., Japan) at 100× magnification. The number of migrated cells was quantified using ImageJ software.

Total protein was extracted from aortic tissues (50 mg, minced on ice) and VSMCs using pre-cooled RIPA lysis buffer (P0013B, Beyotime, Shanghai, China) supplemented with 1× protease/phosphatase inhibitor cocktails. Tissue samples were homogenized (TissueLyser II, QIAGEN N.V., Venlo, The Netherlands) at 50 Hz for 2 min (twice, 30 s interval on ice) with 500 μL lysis buffer; VSMCs were rinsed twice with pre-cooled PBS, scraped in 200 μL lysis buffer per 6 wells, and vortexed gently for 10 min. All lysates were incubated on ice for 30 min and centrifuged at 12,000× g for 15 min (4 °C), and supernatants were collected. Protein concentration was determined via BCA assay (#23225, Thermo, Waltham, MA, USA). Equal protein amounts were separated by 10% SDS-PAGE, transferred to PVDF membranes (IPVH00010, Millipore, Darmstadt, Germany), blocked with 5% non-fat milk in TBST (1 h, RT), incubated with primary antibodies (1:1000, 4 °C, overnight) and HRP-conjugated secondary antibodies (1:5000, 1 h, RT), and visualized with ECL kit (Thermo Fisher, 34580) using an Ultra-Sensitive Multifunctional Imager (Cytiva, Marlborough, MA, USA) to measure the exposure, and band intensities quantified via ImageJ.

Protein samples were collected from treated cells, and protein concentrations were measured using the BCA Protein Assay kit (Pierce^TM, #23225, Thermo, Waltham, MA, USA). Subsequently, 800 μg of protein in lysis buffer was immunoprecipitated using Protein A/G magnetic beads (DynaGreen^TM, Thermo, Waltham, MA, USA). Equal volumes of each protein sample were separated by SDS-PAGE and transferred to PVDF membranes. After blocking with 5% (w/v) non-fat dry milk in TBST for 1 h at room temperature, the membranes were incubated with primary antibody overnight at 4 °C, followed by incubation with HRP-conjugated secondary antibody for 1 h at 25 °C. Protein signals were detected using a chemiluminescence imager [22].

Briefly, purified GST-tagged bait protein and His-tagged prey protein were incubated with glutathione-sepharose beads (Pierce^TM, Thermo, Waltham, MA, USA) in binding buffer (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 1 mM EDTA, 0.1% NP-40, 10% glycerol) at 4 °C for 2 h with gentle rotation. Beads were then washed five times with ice-cold wash buffer to remove non-specific bindings. Bound proteins were eluted by boiling in 2× SDS loading buffer and analyzed by SDS-PAGE followed by Western blotting using anti-His and anti-GST antibodies (abcam). Control groups with GST protein alone were included to confirm specific interactions.

VSMCs in logarithmic growth phase were seeded into 6-well plates at a density of 5 × 10⁵ cells/well and allowed to adhere for 24 h. Cells were then treated according to experimental groups with ox-LDL (50 μg/mL), Ferrostatin-1 (Fer-1, 1 μmol/L), Erastin (10 μmol/L), or their combinations, consistent with Section 2.7. After treatment, cells were washed with pre-cooled PBS, harvested by trypsinization, and centrifuged (1000× g, 5 min, 4 °C). The pellets were resuspended in 200 μL pre-cooled saline, homogenized on ice (30 s × 3 cycles), and centrifuged (3000× g, 10 min, 4 °C) to obtain supernatants. GSH, SOD, MDA, and Fe contents were determined using specific commercial kits (Jiancheng Bioengineering: A006-2-1, A001-3-2, A003-1-2; Abcam: ab83366, Nanjing, China) following manufacturers’ protocols, with three technical replicates and results normalized to protein concentration.

One millimeter of fresh mouse aortic tissue was harvested and quickly frozen on ice. It was immediately placed in a 1.5 mL EP tube of electron microscope fixative solution and fixed at 4 °C in the dark for 24 h, then rinsed 3 times with PBS for 15 min each time. After fixation with osmium tetroxide for 2 h, the cells were rinsed 3 times with PBS for 15 min each time. They were then routinely dehydrated and embedded, stained with lead citrate and uranyl acetate (70–90 nm), and finally imaged with mitochondrial ultrastructure using transmission electron microscopy.

VSMCs were seeded in 6-well plates, and the supernatant was discarded and washed 3 times with PBS. The prepared 1 μmol/L ROS working solution was added, mixed, and incubated in the cell incubator in the dark for 30 min. The plates were resuspended three times in PBS and analyzed by Flow cytometry, and data were analyzed using FlowJo_v10.8.1.

All the data appeared as mean ± standard deviation, and SPSS19.0 software was used to perform statistical analysis. When the data were normally distributed and the variance was homogeneous, one-way analysis of variance (ANOVA) and Student–Newman–Keuls (SNK) test were used for comparison between groups. A p-value less than 0.05 was considered statistically significant.

We initially established models for gene silencing and overexpression of AS and NCOA4. Western blot analysis confirmed the successful establishment of an aorta-specific LvNCOA4 overexpression and shNCOA4 knockdown model in ApoE^−/− mice (Supplementary Figure S1A). Stereomicroscopy and HE staining further validated the promoting effect of NCOA4 in the AS model (Supplementary Figure S1B–D). Additionally, immunofluorescence analysis demonstrated that NCOA4 knockout significantly reduced the area occupied by VSMCs within AS plaques, whereas NCOA4 overexpression yielded the opposite effect (Figure 1A). This observation was corroborated by Western blot results, which indicated a significant upregulation of proliferation and migration markers PCNA and MMP2 in aortic VSMCs from the model group, alongside a notable downregulation of the contractile protein CNN1. In contrast, these effects were reversed in the shNCOA4 group while being exacerbated in the LvNCOA4 group (Figure 1B). These findings suggest that NCOA4 plays a critical role in promoting AS progression, consistent with previous research reports.

To further elucidate the mechanisms underlying the role of NCOA4 in AS progression, we assessed ferroptosis in VSMCs using TEM. TEM analysis revealed a significant reduction in mitochondrial swelling and cristae disruption in VSMCs with NCOA4 knockout, which are characteristic morphological features of ferroptosis (Figure 1C). Immunofluorescence colocalization demonstrated a significant decrease in the colocalization of α-SMA and GPX4 in the HFD group, an effect that was markedly attenuated by NCOA4 knockdown (Figure 1D, Supplementary Figure S1E). Western blot analysis further revealed that the expression of GPX4 and SLC7A11 was significantly increased in the aortas of NCOA4 knockout mice, whereas NCOA4 overexpression led to a decrease in anti-ferroptotic proteins in mouse aortic tissues (Figure 1E). Moreover, biochemical assays confirmed that NCOA4 knockdown inhibited the levels of ferroptosis markers LPO and MDA while upregulating the levels of the antioxidants GSH and SOD. Conversely, NCOA4 overexpression reversed these effects (Figure 1F–I). In summary, these findings demonstrate that NCOA4 promotes the proliferation and migration of VSMCs in AS mice by inducing ferroptosis.

In vitro, we treated VSMCs with ox-LDL and downregulated NCOA4 expression using small interfering RNA (siRNA) (Supplementary Figure S2A). As illustrated in Figure 2A,B, silencing NCOA4 significantly reversed the abnormal proliferation and migration of VSMCs induced by ox-LDL. To investigate whether the effects of NCOA4 silencing on proliferation and migration were mediated through ferroptosis, we introduced the ferroptosis inhibitor Fer-1 and the ferroptosis inducer Erastin. Consistent results from EdU immunofluorescence staining as well as scratch and transwell assays indicated that Erastin treatment could counteract the effects of NCOA4 knockout on both proliferation and migration. Western blot analysis revealed that both Fer-1 treatment and NCOA4 silencing led to a downregulation of key markers associated with proliferation and migration, specifically PCNA and MMP2; however, these effects were reversed by Erastin treatment (Figure 2C). These findings suggest that silencing NCOA4 may inhibit the aberrant proliferation and migration of VSMCs in vitro by suppressing ferroptosis.

Notably, NCOA4 knockout resulted in a significant upregulation of protein expression for critical ferroptosis regulators SLC7A11 and GPX4—an effect that was antagonized by Erastin treatment (Figure 2D). In assays measuring oxidative stress markers, it was observed that NCOA4 silencing effectively mitigated the ox-LDL-induced reduction in levels of antioxidant enzymes such as SOD and GSH while also inhibiting the accumulation of lipid peroxidation products, including MDA and free iron ions (Supplementary Figure S2B–E). Further validation through fluorescence probe assays combined with flow cytometry demonstrated that NCOA4 knockout significantly reduced intracellular ROS levels within VSMCs—a phenomenon abolished upon Erastin treatment (Figure 2E,F). In summary, our results indicate that NCOA4 plays a pivotal role in promoting abnormal proliferation and migration of VSMCs via its mediation of ferroptosis.

Ferritin, composed of FTH heavy chains and FTL light chains, along with FTMT, constitutes essential intracellular iron storage systems [28]. As a selective autophagy receptor for ferritin, NCOA4 serves as a pivotal link between iron metabolism and ferroptosis [29]. To define the role of NCOA4 in iron homeostasis, we established an ox-LDL-stimulated VSMCs model. Notably, the knockout of NCOA4 significantly increased the expression levels of iron storage proteins FTH1 and FTMT without altering FTL levels (Figure 3A,B), consistent with previous reports. FTMT exhibits functional and structural homology with FTH, suggesting conserved roles in regulating the iron–redox balance. Quantitative immunoblotting confirmed that NCOA4 deficiency significantly reduced FTMT expression (Figure 3C,D), implicating NCOA4 in mitochondrial iron regulation. Molecular docking identified a stable NCOA4-FTMT complex stabilized by hydrogen bonds between key residues (Figure 3E). Critically, endogenous Co-IP validated specific NCOA4-FTMT interactions in VSMCs (Figure 3F), while GST pull-down assays using purified GST-NCOA4 demonstrated direct binding (Figure 3G). Immunofluorescence corroborated cytoplasmic colocalization of NCOA4 and FTMT (Figure 3H). These data establish that NCOA4 and FTMT physically interact in VSMCs under physiological conditions.

To further elucidate the regulatory role of this interaction, we employed RNA interference technology to specifically silence the expression of the FTMT gene in VSMCs stimulated by ox-LDL (Supplementary Figure S3). EdU staining analysis demonstrated that silencing FTMT significantly inhibited ox-LDL-induced proliferation of VSMCs (Figure 4A). Transwell migration assays and scratch wound healing experiments indicated that si-FTMT suppressed the enhanced migratory capacity induced by ox-LDL (Figure 4B), accompanied by a reduction in the expression levels of the proliferation marker PCNA and migration-related protein MMP2, while there was an upregulation of contraction phenotype marker CNN1 (Figure 4D). Notably, FTMT silencing markedly decreased intracellular reactive oxygen species (ROS) accumulation (Figure 4C) and reversed ferroptosis driven by ox-LDL, as evidenced by the restoration of GPX4 and SLC7A11 levels (Figure 4D). In summary, these data establish a crucial role for the NCOA4-FTMT interaction in coordinating phenotypic transitions in vascular smooth muscle cells. This is achieved through simultaneous regulation of iron storage dynamics via stabilization of FTMT and modulation of ferroptosis signaling pathways through GPX4/ACSL4 regulation. These findings provide mechanistic insights into how mitochondrial iron homeostasis governs vascular remodeling under conditions of oxidative stress.

To further verify the role of NCOA4 in regulating mitochondrial damage through FTMT-mediated regulation of iron homeostasis within mitochondria, we established FTMT knockdown VSMCs and assessed the subsequent effects on mitochondrial function. The change in mitochondrial membrane potential showed that silencing NCOA4 increased the content of JC-1 aggregates and decreased the content of JC-1 monomer in VSMCs compared with the ox-LDL group, indicating that silencing NCOA4 alleviated ox-LDL-induced mitochondrial membrane potential reduction (Figure 5A). Furthermore, immunofluorescence staining showed that silencing NCOA4 significantly reduced the increase in Mito-Fe²⁺ content in VSMCs induced by ox-LDL. However, we observed that silencing FTMT reversed the therapeutic effects of silencing NCOA4. Dysfunctional iron homeostasis in mitochondria is an important cause of ROS accumulation in mitochondria [30]. Therefore, fluorescence microscopy was used to further investigate the ROS production in mitochondria. We found that silencing NCOA4 again reversed the ox-LDL-induced increase in MitoROS (Figure 5B). Similarly, the content of MitoROS in VSMCs increased again after silencing FTMT (Figure 5C). Taken together, these findings strongly suggest that silencing NCOA4 can regulate mitochondrial iron homeostasis by upregulating FTMT, thereby inhibiting the production of mitochondrial ROS and improving mitochondrial damage.

Mitochondria are the primary site of iron utilization and a key regulator of oxidative metabolism, serving as the principal source of ROS and a crucial locus for ferroptosis [31]. To explore whether silencing NCOA4 inhibits ferroptosis in VSMCs by improving mitochondrial damage, we examined the effect of NCOA4 on mitochondrial integrity and the impact of modulating mitochondrial ROS on ferroptosis. MitoQ, a mitochondrial ROS scavenger, and MitoVES, a mitochondrial ROS inducer, were employed to manipulate MitoROS levels [32,33]. As shown in Figure 6A, silencing NCOA4 significantly reduced ox-LDL-induced mitochondrial damage in VSMCs. In addition, MitoQ was used to inhibit MitoROS production, which could alleviate the abnormal reduction in mitochondrial membrane potential, whereas MitoVES treatment reversed the silencing of NCOA4, consequently exacerbating the impairment of mitochondrial membrane potential. These results indicated that silencing NCOA4 could alleviate mitochondrial damage by inhibiting MitoROS production.

To investigate the role of NCOA4 in modulating VSMCs ferroptosis via mitochondrial damage, we used MitoROS scavengers and inducers to modulate mitochondrial damage and examined their effects on ferroptosis levels in VSMCs. MitoQ was used to inhibit mitochondrial damage, and it reversed ox-LDL-induced decreases in GSH and SOD levels and increases in MDA and LPO levels in VSMCs (Figure 6B–E). Flow cytometry was used to detect the level of ROS in VSMCs, which showed that inhibition of mitochondrial damage reversed the ox-LDL-induced increase in ROS content in VSMCs. (Figure 6F,G). In addition, inhibition of mitochondrial damage significantly increased the protein expression of SLC7A11 and GPX4 compared with the ox-LDL group (Figure 6H). However, the opposite result was observed when MitoVES treatment aggravated mitochondrial damage and MitoVES reversed the therapeutic effect of silencing NCOA4. These findings indicate that mitochondrial damage is positively correlated with ferroptosis in VSMCs. Furthermore, silencing NCOA4 was confirmed to inhibit ferroptosis in VSMCs by reducing mitochondrial damage.

Next, we explored whether NCOA4-induced mitochondrial damage would stimulate the proliferation and migration of VSMCs. We used EdU assays, scratch assays, and transwell assays to assess VSMCs’ proliferation and migration. As shown in Figure 7A,B, treatment with MitoQ significantly reduced EdU fluorescence levels, relative migration rates, and the number of migrating VSMCs compared to the ox-LDL group. Consistent with these results, MitoQ down-regulated the protein expression of PCNA and MMP2 while upregulating CNN1 expression (Figure 7C), indicating that alleviating mitochondrial damage can mitigate abnormal proliferation and migration in VSMCs. Conversely, treatment of VSMCs with MitoVES produced opposite effects. MitoVES induces mitochondrial damage and reverses the inhibitory effect of silencing NCOA4 on abnormal proliferation and migration of VSMCs. Collectively, these results demonstrate that silencing NCOA4 can alleviate ferroptosis and improve VSMCs proliferation and migration by inhibiting mitochondrial damage.

GLXB has a well-documented history in treating AS-related diseases, and our research team confirmed its strong anti-atherosclerotic effects [24]. Assessment of aortic plaque formation through enface photography and HE staining showed that GLXB significantly reduced lipid accumulation and the area of atherosclerotic lesions in AS mice compared to the HFD group in a dose-dependent manner (Figure 8A–C). To further explore GLXB’s impact on VSMCs proliferation and migration within the AS model, we conducted in vitro studies. EdU incorporation assays, wound healing assays, and transwell migration assays revealed that GLXB treatment increased EdU-positive cells compared to the ox-LDL group—indicating reduced inhibition of proliferation—and significantly suppressed both VSMCs migration rate and number (Figure 8D–H). Consistent with these findings, Western blot analysis indicated that GLXB downregulated proliferation marker PCNA and migration-associated protein MMP2 while upregulating contractile phenotype marker CNN1 relative to the ox-LDL group (Figure 8I).

To investigate the role of NCOA4 in the mechanism of GLXB, we assessed its expression levels in aortic plaques. Immunofluorescence staining revealed that NCOA4 levels were significantly elevated in the HFD group compared to the control group; importantly, GLXB treatment was found to dose-dependently suppress this increase (Figure 8J,K). Given the established link between NCOA4 and ferroptosis, we hypothesized that GLXB’s inhibition of plaque formation might involve modulating ferroptosis in VSMCs. To assess ferroptosis within aortic lesions, we performed co-staining for the VSMCs marker α-SMA and the key ferroptosis inhibitor GPX4. This analysis showed that GLXB significantly increased GPX4 protein expression compared to the HFD group (Supplementary Figure S4). Further supporting this, biochemical assays revealed that GLXB treatment effectively lowered the excessive accumulation of MDA and LPO in the aortas of AS mice, while simultaneously boosting levels of reduced GSH and SOD activity (Figure 8L–O). Consistent with these findings, TEM demonstrated that GLXB treatment improved mitochondrial integrity in aortic VSMCs. Specifically, GLXB reduced mitochondrial cristae rupture, decreased the number of vacuolated mitochondria, and attenuated abnormal mitochondrial swelling and shrinkage, resulting in a restoration of near-normal mitochondrial ultrastructure (Figure 8P).

Our previous study demonstrated that GLXB attenuates AS by suppressing LOX-1-dependent cGAS-STING signaling, thereby inhibiting NCOA4-mediated ferroptosis in aortic tissue [22]. To investigate whether the LOX-1 receptor mediates GLXB’s suppression of VSMCs proliferation and migration, we first assessed its protein expression. As illustrated in Supplementary Figure S5, ox-LDL stimulation significantly upregulated LOX-1 expression; however, treatment with GLXB dose-dependently reversed this induction. Subsequently, we examined the functional role of LOX-1 in mediating the effects of GLXB. EdU incorporation assays, scratch wound healing assays, and transwell assays revealed that overexpression of LOX-1 counteracted the inhibitory effects of GLXB on VSMCs proliferation and migration. Specifically, LOX-1 overexpression resulted in an increased area/number of EdU cells (Figure 9A,B) and accelerated wound closure (Figure 9C–E), indicating exacerbated dysregulation of VSMCs.

To assess mitochondrial involvement, we performed JC-1 staining. The results indicated that GLXB significantly mitigated ox-LDL-induced hyperpolarization of mitochondrial membrane potential (ΔΨm) (Figure 9F,H), suggesting protective effects against mitochondrial dysfunction. To establish a connection to ferroptosis, we measured key biomarkers associated with oxidative stress and lipid peroxidation. Notably, GLXB reversed ox-LDL-induced depletion of SOD and GSH while also suppressing accumulations of MDA and total iron levels in VSMCs (Figure 9J–M). Consistent with these findings, treatment with GLXB normalized elevated ROS levels induced by ox-LDL (Figure 9G,I). Mechanistically, Western blot analysis confirmed that GLXB upregulated proteins involved in ferroptosis defense mechanisms—specifically SLC7A11 and GPX4—while downregulating NCOA4 expression. Importantly, these protective effects were abolished upon overexpression of LOX-1 (Figure 9N,O). In summary, our data collectively demonstrate that GLXB mitigates mitochondrial damage and lipid peroxidation within VSMCs through targeting LOX-1; this action inhibits ferroptosis and exerts anti-atherosclerotic effects.