RhoA accelerates atherosclerosis progression by interacting with Hspa5

Abstract

RhoA has been demonstrated to play a role in atherosclerosis (AS); however, its regulatory mechanisms remain poorly characterized. This study aimed to investigate the function of RhoA in AS and elucidate its underlying mechanisms. An AS mouse model was established via a high-fat diet, and the role of RhoA was assessed by administering adeno-associated viruses. Mouse aortic vascular smooth muscle cells (MOVAS) were exposed to oxidized low-density lipoprotein (ox-LDL), and cellular phenotypes were analyzed using cell counting kit-8 assays, Transwell migration assays, transmission electron microscopy, and western blotting. The interaction between RhoA and Hspa5 was investigated through bioinformatic analysis, co-immunoprecipitation, and immunofluorescence. Results revealed that RhoA was highly upregulated in AS mice and predominantly localized in the smooth muscle layer of the aortic root. Knockdown of RhoA or inhibit its activity suppressed the viability, migration, invasion, and mitophagy of ox-LDL-treated MOVAS cells in vitro, and reduced plaque formation and inflammatory responses in AS mice. Furthermore, Hspa5 was found to interact with RhoA, with its expression positively correlated to RhoA levels. Overexpression of Hspa5 counteracted the inhibitory effects of RhoA silencing on cellular behaviors in ox-LDL-stimulated cells in vitro and plaque lesions and inflammation responses in vivo. Collectively, RhoA promotes vascular smooth muscle cell migration, invasion, and mitophagy via interaction with Hspa5, thereby exacerbating AS progression. These findings highlight RhoA as a potential therapeutic target for AS treatment.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-025-33741-w.

Introduction

Atherosclerosis (AS) is a chronic inflammatory vascular disease and a leading cause of global mortality¹. Characterized by abnormal lipid deposition in arterial walls, AS progresses through plaque formation, and unstable plaque rupture ultimately results in arterial narrowing and occlusion². This condition leads to severe complications such as stroke, myocardial infarction, and associated comorbidities that impact quality of life, including depression and arthritis³. Although lipid-lowering therapies have demonstrated efficacy in AS prevention and management, significant clinical challenges persist in its treatment⁴. Therefore, there is an urgent need to elucidate the pathogenic mechanisms of AS to develop effective therapeutic strategies and molecular targets.

Vascular smooth muscle cells (VSMCs) play a pivotal role in the development and progression of AS. Their involvement is multifaceted, including processes like proliferation, migration, and phenotypic switching, which drive plaque progression and instability^5,6. Additionally, interactions between VSMCs and other cell types, such as endothelial cells, critically contribute to vascular calcification and vessel wall remodeling⁷. Hence, understanding VSMC behavior and regulatory mechanisms is critically important for advancing AS research. Targeting VSMC dynamics may yield novel therapeutic approaches to mitigate the cardiovascular burden of atherosclerosis.

Rho family GTPases are key regulators of cytoskeletal dynamics, and their association with AS progression and therapeutic relevance has been extensively documented⁸. RhoA, a member of the Rho family of small GTPases, plays a critical role in modulating cellular processes such as cytoskeletal organization, cell migration, proliferation, and survival⁹. RhoA is highly expressed in vascular endothelium and smooth muscle cells, where it contributes to the regulation of vascular tone through vasoconstriction and vasodilation. The role of RhoA in VSMCs is complex and context-dependent. Under physiological conditions, RhoA signaling is essential for maintaining the differentiated, contractile phenotype of VSMCs, which is crucial for vascular homeostasis¹⁰. However, in response to pathological stimuli such as inflammation and oxidative stress, RhoA activity can drive a phenotypic switch towards a synthetic, proliferative, and migratory state, thereby contributing to vascular remodeling and diseases¹¹. Consequently, RhoA is implicated in various cardiovascular pathologies, including pulmonary arterial hypertension, heart failure, and AS¹². RhoA influences AS progression by modulating the biological activities of multiple cell types, including macrophages, endothelial cells, and VSMCs^13–15. However, the precise mechanisms by which RhoA regulates VSMC behavior remain incompletely understood and warrant further comprehensive investigation.

In the present study, bioinformatics analysis predicted elevated RhoA expression in AS. To validate this hypothesis, we investigated the functional role of RhoA in AS using both in vivo and in vitro experimental models. Additionally, we elucidated the potential molecular pathways underlying RhoA’s regulatory effects. This study provides a more comprehensive theoretical framework for understanding RhoA’s involvement in the pathogenesis of AS.

Materials and methods

Bioinformatic analysis

The GSE215969 dataset was obtained from the Gene Expression Omnibus (GEO) public repository (https://www.ncbi.nlm.nih.gov/geo/). This microarray dataset encompasses mRNA expression profiles from aortic tissues of healthy control mice and AS (ApoE-/-) mice, all maintained on a C57BL/6J genetic background. Differentially expressed genes (DEGs) were identified using R software and defined by thresholds of |log2(fold change)| > 1 and P < 0.05.

Kyoto Encyclopedia of Genes and Genomes (KEGG) enrichment analysis was used to analyze the pathways related to the upregulated genes in AS mice using the DAVID online tool (https://davidbioinformatics.nih.gov/).

Pearson’s correlation coefficient was applied to assess gene co-expression patterns within the RhoA/ROCK signaling pathway.

AS mouse model establishment

Male 8-week-old ApoE -/- mice on a C57BL/6J background were purchased from Biomodel Animal Co., Ltd. (Nantong, China). All animals were maintained in a specific pathogen-free facility under controlled conditions (temperature: 21–23 °C, humidity: 50–70%, 12-hour light/dark cycle). Experimental procedures adhered to the Guide for the Care and Use of Laboratory Animals (8th edition, National Academies Press).

To induce AS, mice were fed a high-fat diet (21% fat, 0.2% cholesterol; Trophic, Nantong, China) commencing at 8 weeks of age for 16 weeks. Control mice received a standard chow diet for the same duration. At the study endpoint, mice were euthanized via CO₂ inhalation followed by cervical dislocation. Blood samples were collected via cardiac puncture, and aortic tissues were perfused with PBS prior to resection. The aorta was severed from the heart just above the aortic synapse, and the aortic root was dissected and preserved for subsequent analysis.

Adeno-associated virus (AAV) injection

Short hairpin RNA targeting RhoA (shRhoA, CAGCTGGACAGGAAGATTATG) and its negative control (shNC, CCTAAGGTTAAGTCGCCCTCG) were designed and cloned into the pAAV-U6-CMV-GFP vectors. The full length of Hspa5 cDNA was cloned into the pAAV-ITR-CMV vector to construct Hspa5 overexpression plasmids. Then, the rAAV2/9-CMV-shRhoA-GFP (AAV-shRhoA), rAAV2/9-CMV-shNC-GFP (AAV-shNC), rAAV2/9-CMV-empty-GFP (AAV-NC), and rAAV2/9-CMV-Hspa5-GFP (AAV-Hspa5) were synthesized. Before AS model generation, the mice were injected with AAVs (1 × 10¹² vg/mouse) through the tail vein as previously described¹⁶.

Oil Red O staining

Aortic root cryosections were prepared by embedding tissues in optimum cutting temperature (OCT) compound (Absin, Shanghai, China). Plaque lesions were assessed using the Oil Red O staining kit (Sbjbio, Nanjing, China). Cryosections (10 μm thickness) were washed with distilled water and incubated in 100% isopropyl alcohol for 5 min. Subsequently, sections were stained with Oil Red O solution at 60 °C for 8 min. Following a 5-minute rinse in 85% isopropyl alcohol and distilled water, Mayer’s hematoxylin solution was applied for nuclear counterstaining (30 s). Finally, stained sections were examined under a light microscope for histological analysis.

En face Oil Red O staining

The whole aorta was opened longitudinally and fixed with 4% paraformaldehyde. The aorta was incubated with Oil Red O solution at room temperature for 15 min, and then washed with 60% isopropyl alcohol for 5 min. The aorta images were taken against a black background.

Enzyme-linked immunosorbent assay (ELISA)

Blood samples were centrifuged at 2000 × g for 10 min, and the resulting serum was collected. The levels of interleukin (IL)-1β, IL-6, and tumor necrosis factor (TNF)-α in the serum were measured using the corresponding mouse ELISA kit (COIBO Bio, Shanghai, China) according to the manufacturer’s instructions.

Cell culture

Mouse aortic vascular smooth muscle cells (MOVAS) were obtained from the American Type Culture Collection (ATCC, Manassas, VA, USA). Cells were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM; Gibco, Grand Island, NY, USA) supplemented with 10% fetal bovine serum (FBS; Procell, Wuhan, China) and 0.2 mg/mL G-418 (Procell) under a humidified atmosphere of 95% air and 5% CO₂ at 37 °C.

To establish an in vitro model mimicking the vascular smooth muscle cell state in AS conditions, MOVAS were treated with 100 µg/mL oxidized low-density lipoprotein (ox-LDL) for 24 h¹⁷. Subsequently, cells were harvested for phenotypic analysis.

Immunofluorescence (IF)

The cross-section of the aorta and aortic root cryosections were fixed in 4% paraformaldehyde and permeabilized with 0.5% Triton X-100. Following blocking with 5% bovine serum albumin (BSA; MCE, Monmouth Junction, NJ, USA), sections were incubated with primary antibodies against SM-MHC (60222-1-Ig and 21404-1-AP, Proteintech, Wuhan, China), RhoA (ab187027, Abcam, Cambridge, MA, USA), and collagen (ab34710, Abcam) overnight at 4 °C. Subsequently, sections were incubated with fluorescently labeled secondary antibodies (goat anti-mouse IgG: ab150113, goat anti-rabbit IgG: ab150078, both from Abcam) at room temperature for 2 h. After counterstaining with 4’,6-diamidino-2-phenylindole (DAPI; MCE) to visualize nuclei, the sections were examined under a confocal laser scanning microscope.

Rhotekin pull-down assay

The aortic root tissues were lysed using the magnesium lysis buffer as previously described¹⁸, and the lysate was collected after centrifuging at 12,000 g for 5 min. The lysate was incubated with Rhotekin-Rho Binding Domain (RBD) agarose beads (Cell Biolabs, San Diego, CA, USA) at 4 °C for 1 h. The agarose beads were washed three times with the lysis buffer and resuspended in Laemmli sample buffer. Them, western blotting was performed to detect RhoA-GTP using the specific primary antibody against RhoA.

Cell transfection

MOVAS cells in the logarithmic growth phase were seeded into 6-well plates. shRhoA, shHspa5, shNC, Hspa5 overexpression plasmids, and empty vectors (pcDNA3.1) were synthesized by Sangon (Shanghai, China). The dominant negative mutant of RhoA (pRK5-myc-RhoA-T19N) was peuchased from Addgene (Cambridge, MA, USA), and the scaffold vector (pRK5-myc; Addgene) was as the negative control. These plasmids were transfected into MOVAS cells using Lipofectamine 2000 reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s protocol.

Quantitative real-time polymerase chain reaction (qPCR)

Total RNA was extracted from cells using TRIzol reagent (Invitrogen). Subsequently, reverse transcription and qPCR were performed with the SYBR One Step RT-qPCR Kit (KeyGEN, Nanjing, China) following the manufacturer’s instructions. Gene expression levels were quantified using the 2^−ΔΔCt method, with β-actin serving as the normalization control. The primer sequences for qPCR were as follows: RhoA forward 5’-AGCTTGTGGTAAGACATGCTTG-3’, reverse 5’-GTGTCCCATAAAGCCAACTCTAC-3’; Hspa5 forward 5’-ACTTGGGGACCACCTATTCCT-3’, reverse 5’-ATCGCCAATCAGACGCTCC-3’; MMP9 forward 5’-CTGGACAGCCAGACACTAAAG-3’, reverse 5’-CTCGCGGCAAGTCTTCAGAG-3’; Lp-PLA2 forward 5’-CTTTTCACTGGCAAGACACATCT-3’, reverse 5’-CGACGGGGTACGATCCATTTC-3’; β-actin forward 5’-GGCTGTATTCCCCTCCATCG-3’; reverse 5’-CCAGTTGGTAACAATGCCATGT-3’.

Cell counting kit-8 (CCK-8)

To assess cell viability, 100 µL of cell suspension containing 2000 cells was seeded into 96-well plates and incubated in a humidified incubator for 24, 48, and 72 h. At each designated time point, 100 µL of CCK-8 solution (Sangon) was added to individual wells followed by 4 h of incubation. Absorbance values were measured at 450 nm using a microplate reader.

Transwell assay

Cell migration and invasion were assessed using Transwell assays with Matrigel-free and Matrigel-precoated chambers (24-well format, Corning, Corning, NY, USA). Cells were suspended in serum-free medium and 1 × 10⁴ cells were seeded into the upper chambers, while the lower chambers were supplemented with complete medium. Following incubation in a humidified atmosphere for 24 h, cells that had migrated or invaded through the filter membranes were fixed in 4% paraformaldehyde and stained with 0.1% crystal violet. Stained cells were visualized under a light microscope. The results were quantified using the ImageJ software.

Transmission electron microscopy (TEM)

The cells were washed with PBS and fixed with 2.5% glutaraldehyde at 4 °C for 30 min. Following a second PBS wash, the cells were post-fixed in 1% osmium tetroxide overnight. After dehydration through a graded ethanol series followed by acetone, the samples were embedded in epoxy resin. Ultrathin section (50 nm) were cut, stained with 3% uranyl acetate and lead citrate, and finally examined under a transmission electron microscope.

Western blotting

Total proteins were extracted from cells using the Total Protein Extraction Kit (KeyGEN), and protein concentrations were measured with the BCA Protein Assay Kit (KeyGEN). Equal amounts of protein were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred onto polyvinylidene difluoride membranes. After blocking with 5% non-fat milk, the membranes were incubated with specific primary antibodies overnight at 4 °C, followed by incubation with HRP-conjugated secondary antibody (ab6721, Abcam) for 2 h at room temperature. Protein bands were visualized using high-sensitivity ECL luminescence reagent (Sangon). The protein levels were quantified using the ImageJ software. The primary antibodies were as follows: anti-RhoA (ab187027, Abcam), anti-PINK1 (23274-1-AP, Proteintech), anti-Parkin (ab324566, Abcam), anti-LC3B (ab192890, Abcam), anti-Beclin 1 (ab207612, Abcam), anti-p62 (31403-1-AP, Proteintech), anti-Hspa5 (ab21685, Abcam), and anti-β-actin (ab8227, Abcam).

Co-immunoprecipitation (co-IP)

Endogenous co-IP was performed in MOVAS cells, while exogenous co-IP was conducted in HEK293T cells (ATCC). The Protein A/G magnetic IP/Co-IP kit (Absin) was used in this study. Briefly, HEK293T cells were cultured at DMEM supplemented 10%FBS at 37 °C with 5% CO₂. These cells were lysed in lysis buffer on ice for 5 min, and the lysate was collected. The magnetic beads were washed with wash buffer and incubated with antibodies at room temperature for 2 h. The antibody-labeled beads were incubated with cell lysate at room temperature for 30 min. The magnetic beads were washed with wash buffer, and the target antigen was denatured using electrophoretic loading buffer at 100 °C for 10 min. Western blotting was performed to detect protein levels.

Statistical analysis

Data are expressed as the mean ± standard deviation. Student’s t-test was used to analyze the difference between the two groups, and one-way ANOVA (analysis of variance) was used to compare the differences among multiple groups. P < 0.05 was considered statistically significant. Data analysis was performed using GraphPad Prism 8.

Results

RhoA is highly expressed in the vascular smooth muscle of mice with AS

We first performed microarray analysis to identify DEGs between AS and healthy control mice. The results are shown using a volcano plot (Fig. 1A) and a heatmap (Fig. 1B). A total of 85 upregulated genes and 33 downregulated genes were identified in AS. Among these, RhoA expression was predicted to be upregulated in AS. Several previous studies have demonstrated that RhoA is involved in the progression of AS; however, its role in regulating cell phenotypes and underlying molecular mechanisms remains poorly characterized. Thus, we selected RhoA for further investigation. We then generated an AS mouse model. The results of en face oil red O staining showed that AS lensions could be seen in the aortas of the AS mice, while no obvious AS lesions were observed in the aortas of the control group mice (Fig. 1C). The levels of pro-inflammatory factors IL-1β and IL-6 were significantly higher in the serum of AS mice compared to those in control mice (Fig. 1D,E). These results confirm the successful establishment of the AS mouse model. To explore the role of RhoA in AS, we measured its expression in mice using IF. SM-MHC staining was used to identify smooth muscle cells. We observed that RhoA was predominantly expressed in the smooth muscle of the aortic root, with significantly increased expression in the smooth muscle of the aortic root in AS mice compared with the control group (Fig. 1F). In addition, we measured RhoA activity. As shown in Fig. 1G,H, RhoA activity was elevated in the aortic root of AS mice, compared with the control group. These findings suggest that RhoA may regulate AS progression through modulation of the biological behaviors of smooth muscle cells.

Fig. 1.

RhoA is highly expressed in the vascular smooth muscle of mice with AS. (A) DEGs were predicted using the GSE215969 microarray and shown using a volcano map. Red: upregulated genes; blue: downregulated genes; gray: no significant difference. (B) DEGs from the GSE215969 microarray were also shown using a heat map. Red: upregulation; blue: downregulation. AS mouse model was established using high-fat diet, while the control group mice were fed standard chow. (C) Representive images of en face Oil Red O staining assay. Scale bar = 2 mm. The serum was collected from mice, and ELISA was conducted to measure the levels of (D) IL-1β and (E) IL-6. After collecting the aortic root from mice, (F) the location and expression of RhoA were visualized using IF, and SM-MHC was stained to show the smooth muscle cells. Scale bar = 100 μm. (G) RhoA activity was detected using western blotting ater Rhotekin pull-down assay, and the results were quantified in (H). n = 6/group.

Knockdown of RhoA inhibits MOVAS cell viability, migration, invasion, and mitophagy

To explore the effect of RhoA on the phenotypes of smooth muscle cells, we used MOVAS cells. The expression of RhoA was knocked down by transfection with shRhoA (Fig. 2A). MOVAS cells were stimulated with ox-LDL to create a cell model. Cellular behaviors were evaluated. The results of the CCK-8 assay indicated that ox-LDL treatment enhanced cell viability, whereas RhoA knockdown reduced cell viability in ox-LDL-treated MOVAS cells (Fig. 2B). Transwell assay results demonstrated that ox-LDL promoted the migration and invasion of MOVAS cells, while RhoA silencing counteracted this effect (Fig. 2C–F). Mitophagy was analyzed using TEM. We observed that ox-LDL induced mitophagosome formation and reduced mitochondrial content, which were rescued by RhoA knockdown (Fig. 2G). Furthermore, ox-LDL stimulation upregulated the protein levels of PINK1, Parkin, Beclin-1, and the LC3II/I ratio, while downregulating p62 levels in MOVAS cells. These effects were reversed by RhoA knockdown (Fig. 2H-M). Collectively, these findings demonstrate that RhoA knockdown suppresses ox-LDL-induced increases in cell viability, migration, invasion, and mitophagy in MOVAS cells.

Fig. 2.

Knockdown of RhoA inhibits MOVAS cell viability, migration, invasion, and mitophagy. (A) MOVAS cells were transfected with shNC and shRhoA, and qPCR was conducted to determine RhoA expression. After knocking down RhoA expression, MOVAS cells were treated with ox-LDL, and cell phenotypes were assessed. (B) Cell viability was detected using CCK-8. (C,D) Migration and (E,F) invasion were evaluated using Transwell assay, and their rates were quantified. Scale bar = 100 μm. (G) Mitophagy was observed using TEM. (H) The levels of PINK1, Parkin, LC3 (LC3I/II), Beclin-1, and p62 were measured using western blotting. β-actin served as the internal control. The protein levels of (I) PINK1, (J) Parkin, (K) LC3II/I, (L) Beclin-1, and (M) p62 were quantified. n = 3/group.

Inactivation of RhoA suppresses MOVAS cell viability, migration, invasion, and mitophagy

To further explore the involvement of RhoA in cellular behaviors, we transfected dominant-negative mutant RhoA-T19N and its negative control vectors into MOVAS cells. We found that RhoA-T19N reversed the promotion of cell viability, migration, and invasion caused by ox-LDL (Fig. S1A–E). Moreover, ox-LDL treatment induced the upregulation of PINK1, Parkin, LC3II/LC3I, and Beclin-1 and the downregulation of p62 in MOVAS cells, whereas RhoA-T19N counteracted the effect on the levels of these markers induced by ox-LDL (Fig. S1F–K). The results demonstrate that inhibiting the activation of RhoA suppresses ox-LDL-induced increases in cell viability, migration, invasion, and mitophagy in MOVAS cells.

RhoA interacts with Hspa5

To investigate the molecular mechanism, all upregulated genes in AS identified from the microarray data were subjected to KEGG enrichment analysis. The results revealed that these genes were enriched in multiple pathways, particularly the RhoA/ROCK signaling pathway, in which RhoA plays a central regulatory role (Fig. 3A). Next, the Pearson correlation coefficient was calculated to assess the pairwise correlations among these factors within the pathway. We observed a positive correlation between RhoA and Hspa5 (Fig. 3B). As previously described, Hspa5 plays a role in AS progression by regulating vascular endothelial cell proliferation and apoptosis¹⁹; however, whether Hspa5 regulates VSMC behaviors remains unclear. Thus, Hspa5 was selected for the following study. The protein levels of Hspa5 were analyzed in MOVAS cells, and Western blot results demonstrated that RhoA knockdown significantly reduced Hspa5 expression (Fig. 3C–E). Both endogenous and exogenous co-IP experiments confirmed that RhoA physically interacts with Hspa5 (Fig. 3F,G).

Fig. 3.

RhoA interacts with Hspa5. (A) The pathways related to upregulated genes in AS that were predicted using the GSE215969 microarray were predicted using the KEGG enrichment analysis and shown with a dot bubble diagram. (B) The correlation of the factors in the RhoA/ROCK pathway was analyzed by Pearson correlation coefficient and displayed with a bubble diagram. (C) The levels of Hspa5 in MOVAS cells after knocking down RhoA. (D) RhoA and (E) Hspa5 levels were quantified. (F) Endogenous and (G) exogenous co-IP were performed to verify the interaction between RhoA and Hspa5 proteins. n = 3/group.

Knockdown of Hspa5 inhibits MOVAS cell viability, migration, invasion, and mitophagy

To explore the role of Hspa5 in vitro, shNC and shHspa5 were transfected into MOVAS cells. Following shHspa5 transfection, the expression of Hspa5 was decreased (Fig. S2A). In ox-LDL-induced MOVAS cells, knockdown of Hspa5 suppressed cell viability (Fig. S2B), migration (Fig. S2C,D), invasion (Fig. S2E,F), and mitophagosome formation (Fig. S2G). The levels of PINK1, Parkin, LC3II/LC3I, and Beclin-1 were decreased, while p62 levels were elevated after Hspa5 knockdown in ox-LDL-induced MOVAS cells (Fig. S2H–M). The results demonstrate that knockdown of Hspa5 inhibits ox-LDL-induced viability, migration, invasion, and mitophagy.

Overexpression of Hspa5 reverses the inhibition of cell viability, migration, invasion, and mitophagy induced by RhoA knockdown

To investigate the role of both RhoA and Hspa5 in vitro, MOVAS cells were transfected with Hspa5 overexpression plasmids, while empty vectors served as negative controls. qPCR results demonstrated a significant upregulation of Hspa5 expression following plasmid transfection (Fig. 4A). Subsequently, the functional impact of Hspa5 was further assessed through rescue experiments. MOVAS cells were subjected to RhoA knockdown combined with Hspa5 overexpression and then stimulated with ox-LDL. The results revealed that RhoA silencing significantly suppressed cell viability, migration, invasion, and mitophagy, whereas Hspa5 overexpression effectively reversed these inhibitory effects (Fig. 4B–G). In cells treated with ox-LDL, RhoA knockdown led to decreased levels of PINK1, Parkin, LC3II/I, and Beclin-1, accompanied by increased p62 accumulation. Notably, Hspa5 overexpression completely abrogated these alterations induced by RhoA depletion (Fig. 4H–M). Collectively, these findings demonstrate that RhoA knockdown impairs MOVAS cell viability, migration, invasion, and mitophagy through suppression of Hspa5 expression.

Fig. 4.

Overexpression of Hspa5 reverses the inhibition of cell viability, migration, invasion, and mitophagy induced by RhoA knockdown. (A) MOVAS cells were transfected with empty vector and Hspa5 overexpression plasmids, and qPCR was conducted to determine Hspa5 expression. MOVAS cells were transfected with shRhoA and Hspa5 overexpression plasmids, followed by treatment with ox-LDL, and cell phenotypes were assessed. (B) Cell viability was detected using CCK-8. (C,D) Migration and (E,F) invasion were evaluated using Transwell assay, and their rates were quantified. Scale bar = 100 μm. (G) Mitophagy was observed using TEM. (H) The levels of PINK1, Parkin, LC3 (LC3I/II), Beclin-1, and p62 were measured using western blotting. β-actin served as the internal control. The protein levels of (I) PINK1, (J) Parkin, (K) LC3II/I, (L) Beclin-1, and (M) p62 were quantified. n = 3/group.

Silencing of RhoA attenuates plaque lesions and inflammatory response in AS mice by decreasing Hspa5 expression

To investigate the role of RhoA and Hspa5 in vivo, AAV-shNC, AAV-shRhoA, AAV-NC, and AAV-Hspa5 were administered to AS mice. The results of qPCR showed that Hspa5 expression was elevated in the AS group. In AS mice, knockdown of RhoA decreased Hspa5 expression, which was reversed by AAV-Hspa5 (Fig. 5A). En face Oil Red O staining results showed that knockdown of RhoA attenuated AS lensions in the whole aorta of AS mice, which was reversed after Hspa5 overexpression (Fig. 5B). Plaque lesion formation in the aortic root was assessed via Oil Red O staining, revealing that RhoA knockdown significantly reduced lesion size in atherosclerotic plaques (Fig. 5C,D). IF results showed that the atherosclerotic plaque lesions in the AS group showed lower smooth muscle cells and collagen than that in the control group. Knockdown of RhoA increased smooth muscle cells and collagen, while Hspa5 abrogated the effect caused by RhoA silence (Fig. 5E). MMP9 and Lp-PLA2 are potential biomarker for AS plaque vulnerability^20,21. Thus, we measured their expression to assess AS plaque stability. qPCR results showed that MMP9 and Lp-PLA2 expression was upregulated in the AS group, indicating plaque vulnerability. RhoA knockdown decreased MMP9 and Lp-PLA2 expression, which was reversed by Hspa5 overexpression (Fig. 5F,G), suggesting that RhoA knockdown helps stabilize plaques, while Hspa5 impairs the stability of plaques. Furthermore, RhoA silencing resulted in significantly lower serum concentrations of pro-inflammatory cytokines IL-1β, IL-6, and TNF-α in AS mice (Fig. 5H–J).

Fig. 5.

Silencing of RhoA attenuates plaque lesions and inflammatory response in AS mice. AS mouse model was established using high-fat diet, and AAV-shRhoA, AAV-shNC, AAV-NC, and AAV-shHspa5 were injected through the tail vein. (A) Hspa5 expression was measured using qPCR. The whole aorta was obtained from mice, and (B) representive images of en face Oil Red O staining. Scale bar = 2 mm. The aortic root was obtained from mice, and (C) Oil Red O staining was performed to determine plaque lesions, and (D) the staining area was quantified. Scale bar = 200 μm. (E) Representive images of IF staining to observe smooth muscle cells (SM-MHC staining) and collagen (collagen staining). Scale bar = 50 μm. (F) MMP9 and (G) Lp-PLA2 expression was detected using qPCR. The serum was collected from mice, and the levels of (H) IL-1β, (I) IL-6, and (J) TNF-α were measured by ELISA. n = 6/group.

Discussion

In the present study, we investigated the role of RhoA in AS. An AS mouse model was established, and increased plaque lesions were observed, confirming successful model establishment. Additionally, elevated levels of pro-inflammatory factors in AS mice indicated that inflammation plays a pivotal role in disease progression, aligning with the well-established characterization of AS as a chronic inflammatory disorder^22,23. RhoA expression was significantly upregulated in the aortic smooth muscle of AS mice, and RhoA activity was also elevated. Knockdown of RhoA attenuated plaque lesion formation and reduced inflammatory responses, suggesting that under AS conditions, RhoA predominantly exerts a pathogenic role by modulating VSMC behaviors.

VSMCs are critical drivers of AS pathogenesis. In healthy blood vessels, VSMCs exhibit low proliferation rates; however, in AS, these cells infiltrate atherosclerotic plaques and exacerbate lesion expansion through enhanced migratory and invasive capacities²⁴. Prior studies have demonstrated that RhoA promotes VSMC proliferation, migration, and invasion^10,25. In this study, we utilized ox-LDL-stimulated VSMCs to model AS in vitro, as elevated LDL levels are a well-established risk factor for the disease²⁶. Our results revealed that RhoA knockdown suppressed these pathological behaviors in VSMCs, consistent with its established role in promoting the synthetic, pathological phenotype. While RhoA is necessary for contractile function in a healthy vessel, its sustained activation in the pro-inflammatory and pro-lipidic microenvironment of AS appears to lock VSMCs into this deleterious synthetic state.

Additionally, we observed that RhoA silencing inhibited mitophagy in VSMCs. Mitophagy, a process that clears damaged and dysfunctional mitochondria, plays a critical role in maintaining cardiovascular homeostasis. Its dysregulation has been implicated in AS through effects on vascular endothelial cells, macrophages, and VSMCs^27,28. Notably, VSMC phenotype switching is closely linked to mitochondrial dynamics and mitophagy, which directly regulate VSMC proliferation and migration²⁹. The findings regarding mitophagy in this study represent a novel contribution to the field and align with prior reports demonstrating RhoA-mediated promotion of mitophagy in other pathological contexts^30,31. Our study also indicated that the inactivation of RhoA using its dominant-negative forms (T19N) inhibited VSMC viability, migration, invasion, and mitophagy. These findings indicate that the promoting effect of RhoA on AS not only depends on the changes in its expression level but is also related to its GTP activity.

It is worth noting that the function of RhoA is highly dependent on cell type and disease background. For instance, recent studies have shown that under the pathological conditions of microvascular systems (such as cutaneous arterioles), such as Raynaud’s phenomenon, the activation of RhoA can mediate the transport of α2C-adrenergic receptors to the cell membrane, thereby regulating the vasoconstriction response³². These findings collectively highlight the complexity of the RhoA signaling pathway, whose role in vascular diseases depends on the specific cellular microenvironment and functions.

Subsequent analyses revealed a positive correlation between Hspa5 and RhoA expression, with evidence of functional interaction between these proteins. Hspa5, also known as GRP78 or BIP, is an endoplasmic reticulum-resident chaperone of the Hsp70 family³³. Under physiological conditions, Hspa5 is expressed at low levels³⁴; however, its upregulation is associated with various pathological processes³⁵. Previous studies have highlighted Hspa5’s role in promoting inflammation and endothelial cell proliferation in AS^19,36, yet its regulatory effects on VSMC behavior remain poorly characterized. Elevated Hspa5 expression has been reported in aortic VSMCs of AS mouse models³⁷. In this study, in vitro experiments demonstrated that Hspa5 overexpression counteracted the suppression of VSMC viability, migration, invasion, and mitophagy induced by RhoA knockdown, suggesting a pro-atherogenic role for Hspa5. Moreover, the in vivo experiments showed that overexpression of Hspa5 abrogated the inhibition of AS plaque lesions and inflammation response caused by RhoA knockdown. The results demonstrate that silencing of RhoA attenuates AS by decreasing Hspa5 expression.

Beyond establishing a physical interaction, our data prompt the question of how RhoA and Hspa5 are co-regulated within the atherosclerotic microenvironment. We propose a positive regulatory mechanism that induces VSMC dysfunction. In AS, both the expression of RhoA and Hspa5 was elevated, and RhoA positively regulated Hspa5 expression. We speculate that simultaneously regulating RhoA and Hspa5 may enhance the influence on the behavior of VSMCs, thereby playing an important role in the progression of AS. This newly discovered molecular mechanism is different from its classic function of regulating the cytoskeleton through ROCK. A recent study in the field of hypertension has revealed a novel regulatory paradigm targeting the RhoA signaling pathway³⁸. This study discovered the small molecule compound AH001, which does not directly inhibit the GTPase activity of RhoA, but rather promotes the interaction between RhoA and the inhibitory factor RhoGDI1 by binding to the TRPV4 ion channel. These findings suggest that interventions targeting the upstream regulatory nodes of RhoA or its non-classical interacting proteins may be more specific and feasible than directly targeting RhoA itself, providing key ideas for the development of new strategies for treating vascular diseases.

The therapeutic potential of modulating the RhoA pathway in atherosclerosis is indirectly supported by existing clinical and preclinical evidence. Statins, as the cornerstone of AS treatment, have been found to inhibit RhoA^39,40. Therefore, we propose that RhoA may serve as a therapeutic target for Statins. In future work, we can further verify this hypothesis through animal models to provide better support for the clinical translation of this study.

There are several limitations in this study. First, we used MOVAS cells to perform functional assays. Although our findings reveal a novel mechanistic axis in AS, future validation in human primary VSMCs will be crucial to fully ascertain its clinical relevance. Second, although we found that RhoA activity was elevated in AS mice and inactivation of RhoA inhibits VSMC cell behaviors, whether its role in vivo is related to its activity remains unclear and needs further investigation. Additionally, the animal experiments were conducted exclusively in male mice. While this choice was made to minimize hormonal variability and establish a proof-of-concept, it precludes the extrapolation of our findings to females. Future studies are essential to investigate the role of the RhoA/Hspa5 axis in female animal models and to explore potential sex-specific differences in this signaling pathway.

In conclusion, our study demonstrates that RhoA is significantly upregulated in the aortic smooth muscle of AS mice. Knockdown of RhoA decelerates AS progression by inhibiting VSMC migration, invasion, and mitophagy through interaction with the Hspa5 protein. These findings reveal that the RhoA/Hspa5 axis plays a critical role in AS pathogenesis and identify RhoA as a potential therapeutic target for AS treatment.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

All authors participated in the design, interpretation of the studies and analysis of the data and review of the manuscript. R D drafted the work and revised it critically for important intellectual content; C C, J L, G J and Y T were responsible for the acquisition, analysis and interpretation of data for the work; X S made substantial contributions to the conception or design of the work. All authors read and approved the final manuscript.

Funding

The work was supported by Hebei Natural Science Foundation under grant number H2025307059.

Data availability

The datasets used and/or analysed during the current study are available from the corresponding author on reasonable request.

Declarations

Competing interests

The authors declare no competing interests.

Ethics approval and consent to participate

This study was approved by the Ethics Committee of Hebei General Hospital(Approval No.2024-DW-005). All animal experiments should comply with the ARRIVE guidelines. All methods were carried out in accordance with relevant guidelines and regulations.