Establishment of a medial arterial calcification model in C57BL/6J mice via arterial intimal injury

Introduction

Medial arterial calcification (MAC) is a chronic systemic vascular disease, which is characterized by the diffusion and progressive precipitation of calcium phosphate in the medial artery. Hence, patients with MAC often exhibit increased hardness and decreased compliance of blood vessels, tend to have hemodynamic disorders and chronic severe limb ischemia, and are more susceptible to cardiovascular disease and death. MAC often occurs among older adults and patients with chronic kidney disease (CKD), diabetes mellitus (DM), or hypertension (1,2). Because the initial clinical symptoms are often not obvious and the rate of diagnosis is low, MAC has often been an afterthought in clinical practice and has not received sufficient attention in this context. However, an accumulating body of research indicates that MAC is a key negative predictor of cardiovascular morbidity and mortality and should be considered to be a silent killer of the cardiovascular system (3,4).

The molecular pathogenesis of MAC and related risk factors remains unclear, and animal models are an important means for clarifying them. Rodents are less prone to vascular calcification, and the majority of vascular calcification models trigger the MAC formation by inducing CKD or DM through kidney removal, a high amino acid diet, a high vitamin D₃ (VD₃) diet, a high phosphate diet, or nicotine administration (5,6). These can possibly cause other undesired complications such as acute renal failure or multiple organ calcification, complicating the evaluation of the risk factors related to MAC formation (7-9). Therefore, animal models that enable better detection and assessment of the risk factors of MAC pathogenesis are required.

Vascular calcification, particularly MAC, is increasingly recognized as a process resembling endochondral ossification. This pathological transition is primarily driven by chondrocyte-like differentiation of vascular smooth muscle cells (VSMCs), rather than direct osteoblast-like transformation. Under stimuli such as inflammation, oxidative stress, or shear pressure, VSMCs undergo phenotypic switching toward a chondrogenic lineage, characterized by expression of SRY-box transcription factor 9 (Sox9) and secretion of cartilage-specific extracellular matrix proteins such as type II collagen and aggrecan. This cartilage-like template subsequently becomes mineralized, leading to vessel wall calcification (10,11). This mechanism has been consistently observed in both experimental models and human studies of MAC. The mouse carotid artery wire injury model is a well-established tool for studying endothelial regeneration (12). While severe endothelial dysfunction is a known precursor to vascular calcification, reliably modeling MAC has remained challenging. Driven by our initial serendipitous observation that varying the degree of mechanical injury (by adjusting guidewire diameter and friction) could induce varying degrees of vascular calcification, we sought to systematically optimize the model. We then rigorously evaluated key parameters—including guidewire diameter, the timing of calcification development, and the role of VD₃ supplementation—to establish a definitive and optimal protocol for inducing MAC in C57BL/6J mice. Using this model, we not only preliminarily established the pathogenic standards for MAC but also examined its response to a calcification inhibitor, thereby validating its utility for therapeutic screening. We present this article in accordance with the ARRIVE reporting checklist (available at https://cdt.amegroups.com/article/view/10.21037/cdt-2025-435/rc).

Methods

Animals

Seven- to eight-week-old male SPF C57BL/6J wild-type mice were purchased from GemPharmatech Co., Ltd. (Nanjing, China). All mice were housed under a 12-hour light/dark cycle in a controlled environment (23±2 ℃, 50%±10% humidity, 3–5 mice per cage). Health monitoring was performed weekly, and all mice remained healthy throughout the acclimatization period.

All animal experiments were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. The study protocols were approved by the Animal Welfare and Ethics Committee of Huzhou University (No. 20220701). A protocol was prepared before the study without registration. Mice were humanely euthanized via CO₂ asphyxiation at the desired time points. Every effort was made to minimize suffering. Humane endpoints were strictly enforced to prevent undue suffering. No animals (humane endpoints: >20% weight loss, severe lethargy/unresponsiveness, inability to eat/drink, or signs of severe infection) met these criteria, and no adverse events or mortality occurred.

Carotid artery injury and MAC model construction

After a 1-week acclimation period, 2-month-old mice were fed a chow diet and provided with water ad libitum. Mice were randomly assigned to experimental groups using a random number generator: control group, wire diameter groups, modeling time groups, and intervention groups. Figure 1A shows the experimental groups and the number of animals (n) in each group. For the surgery, anesthesia was induced via isoflurane (Catalog #R510-22-16, RWD, Shenzhen, China) at 5% and 2% for induction and maintenance, respectively, and carprofen was injected subcutaneously for analgesia (5 mg/kg) (Catalog #HY-B1227, MedChem Express, Monmouth Junction, USA). The depth of anesthesia was confirmed by the absence of pedal reflex. The epilated area was disinfected with iodophor, and a midline incision (8–10 mm) was made in the neck of the mouse after anesthesia. The left carotid artery (LCA), external carotid artery (ECA), and internal carotid artery (ICA) were exposed via blunt dissection. The LCA and ICA were occluded temporarily via microsurgical arterial clips, while the ECA was ligated with 6-0 silk suture. A minimal incision was created in the ECA using microscissors. Subsequently, a polished stainless steel wire was inserted 5 mm into the LCA through the incision. This procedure was repeated three times to induce consistent endothelial injury. Three distinct diameters of stainless steel wires were employed across three experimental groups, with all wires subjected to standardized surface roughening via microgrinding to ensure consistent texture irrespective of diameter. The microsurgical arterial clips were then released to restore blood flow, leaving blood to flow through the ICA via the LCA. The subcutaneous tissue was carefully repositioned, and the epidermal layer was subsequently sutured. For the sham operation group, temporary occlusion of the LCA and ICA, mini-incision, and ligation of the ECA were performed, but mechanical injury with wire was not. The operation was completed under a binocular stereo microscope. The surgical procedure lasted approximately 30 minutes. Mice were monitored following surgery until recovery while they were housed in a chamber on a heating pad. For the VD₃ experimental group, each mouse was administered VD₃ subcutaneously (500,000 IU/kg) (CAS 67-97-0, Catalog #C1234, Yuanye Bio, Shanghai, China) for 3 consecutive days beginning the second day after mechanical injury to the LCA. VD₃ was prepared according to Kwon et al. (13) as follows: 14.575 mg of VD₃ was first dissolved in 70 µL of anhydrous ethanol, then mixed with 500 µL of castor oil at room temperature (suspended for 15 minutes for complete dissolution), and finally mixed with 6.2 mL of sterile injection water.

Figure 1 Group conditions and MAC mouse model construction. (A) Overall study design and workflow. (B) Schematic diagram of the CCA endothelial injury. The boxed area is shown as an enlarged view. (C) Macroscopic comparison of CCA morphology before and after mechanical endothelial injury. CCAs in the sham-operated group exhibited a smooth surface and uniform diameter. In contrast, CCAs in the experimental group appeared dilated, with an irregular diameter and localized whitish areas (indicative of injury). (D) Representative H&E staining of the blood vessels of the control and experimental groups. CCA, common carotid artery; H&E, hematoxylin and eosin; MAC, medial arterial calcification; VD₃, vitamin D₃.

Following the successful establishment of the MAC model, therapeutic interventions were initiated to evaluate the efficacy of different anti-calcification regimens. In one experimental group, beginning 1 week after surgery, each mouse received 180 mg/kg of the calcification inhibitor etidronate acid disodium (CAS 7414-83-7, Catalog #D135681, Alladine, Shanghai, China) daily via oral gavage for 2 consecutive weeks. In another group, starting from the second day after surgery, each mouse was administered 15 mg/kg of the calcification inhibitor SNF472 (Catalog #HY-NO814A, MedChem Express) subcutaneously daily. This regimen consisted of three treatment courses over 12 days (14).

To minimize potential confounding factors, the sequence of animal surgery, drug administration, tissue harvesting, and histological processing was randomized throughout the study. Furthermore, blinding was implemented during group allocation, all surgical and therapeutic procedures, histological staining, and data analysis. Consequently, all animals subjected to the intervention were included in the final analysis, with no data points or animals excluded.

Description of calcification inhibitors

Etidronate, a bisphosphonate, inhibits ectopic calcification by directly suppressing hydroxyapatite crystal formation and growth. It exhibits high affinity for calcium phosphate, adsorbing onto developing crystal surfaces to prevent further maturation and aggregation. Clinically approved for conditions such as heterotopic ossification, Paget’s disease and Generalized Arterial Calcification of Infancy (15), its efficacy in inhibiting vascular calcification has been demonstrated in both preclinical and clinical studies.

SNF472, a sodium salt of phytic acid derived from plants, selectively targets and inhibits hydroxyapatite crystallization. It specifically binds to active calcification sites, stabilizing amorphous calcium phosphate precursors and preventing their transformation into pathological hydroxyapatite crystals. Currently under investigation as an innovative therapy for vascular calcification, SNF472 has shown promising results in clinical trials involving hemodialysis patients with end-stage renal disease, effectively delaying the progression of cardiovascular calcification (14).

Hematoxylin and eosin (H&E) and Von Kossa staining

When the modeling time (1, 2, 3, and 4 months) was reached, the length of 4–5 mm injury sites in the LCA was extracted. The blood vessel samples were fixed with paraformaldehyde for H&E and Von Kossa staining. None of the tissue sections underwent decalcification. All stained samples were scanned with a panoramic digital scanner (KF-PRO-120-HI, Jiangfeng Bio, Shanghai, China) using the manufacturer’s accompanying KF-PRO-SCAN software (version 1.7.1.1), and photographed with a 40× objective lens. Analysis of the images was performed with the corresponding digital imaging software. All the samples were numbered, and the pathologist performed a blinded analysis of the results, and their evaluations were averaged to determine the final grade for each sample.

Immunohistochemical staining

Blood vessel samples were embedded in paraffin and sectioned at 5 µm thickness for immunohistochemistry (IHC). After baking at 62 ℃, dewaxing, and hydration, endogenous peroxidase activity was blocked with 3% H₂O₂. Tris-EDTA buffer was used for heat-mediated antigen retrieval. Subsequently, sections were blocked with 10% normal goat serum in 1% bovine serum album (BSA) for 1 hour at room temperature to minimize non-specific binding. After blocking, sections were incubated overnight at 4 ℃ with the following primary antibodies diluted in 1% BSA: anti-runt-related transcription factor 2 (Runx2) (1:100; ab192256; Abcam, Cambridge, UK), anti-alpha smooth muscle actin (α-SMA) (1:4,000; ab7817; Abcam), anti-osteopontin (OPN) (1:200; 25715-1-AP; Proteintech, Chicago, USA), anti-interleukin-6 (IL-6) (1:100; ab290735; Abcam), or anti-interleukin-1beta (IL-1β) (1:200; ab283818; Abcam). The following day, corresponding secondary antibody and diaminobenzidine (DAB) were applied for incubation at 37 ℃. Cell nuclei were stained via the EnVision method.

Immunofluorescence (IF)

Blood vessel samples (7 µm) were paraffinized for IF staining. After dewaxing and antigen retrieval, sections were blocked with 10% normal goat serum and 1% BSA in phosphate-buffered saline (PBS) for 1 hour at room temperature. Subsequently, samples were incubated with rabbit anti-CD86 polyclonal antibody (1:100; Cat #19589; Cell Signaling Technology, Danvers, USA) diluted in 1% BSA/PBS at 4 ℃ overnight. After PBS washes, samples were incubated with Alexa Fluor 488-conjugated goat anti-rabbit IgG (1:2,000; Cat #ab205718; Abcam) for 1 h at room temperature. Cell nuclei were stained with 4',6-diamidino-2-phenylindole (DAPI) (1 µg/mL) for 10 min at room temperature. Fluorescent images were captured using a DS-Fi2 camera (Nikon) with consistent exposure settings for all samples.

The histopathological grading system for mouse MAC

By using clinical classification criteria for vascular calcification as a reference (16-18), we developed a histopathological grading system for mouse MAC according to the length of the calcium deposition observed with H&E and Von Kossa staining.

Statistics analyses

This study established a foundational characterization of the calcification model using semi-quantitative grading. All data were analyzed with SPSS software (IBM Corp., Armonk, NY, USA). For comparisons involving ordinal rating data, Fisher’s exact probability test was applied, as appropriate. A P value <0.05 was considered statistically significant.

Results

Establishment of the MAC mouse model

As shown in Figure 1B-1D, compared to the sham group, which exhibited normal arterial structure, the experimental group subjected to mechanical injury of the common carotid artery (CCA) using 0.45 mm rough stainless steel guide wires developed significant vascular calcification after 3 months, as evidenced by reduced arterial compliance, increased stiffness, and marked thickening of the intima and media.

Histological examination revealed that VSMCs in the experimental group underwent activation, proliferation, and migration into the intima, where they differentiated into cells exhibiting a chondroid morphology, with small cell volumes, scattered within the CCA (Figure 2A-2C). These cells further proliferated and were associated with the deposition of a basophilic, cartilage-like matrix (Figure 2D,2E). A subset of these cells hypertrophied and formed homogeneous cell clusters, resembling those seen in hyaline cartilage (Figure 2F), while other areas exhibited features indicative of osteogenic differentiation (Figure 2G,2H). IF analysis confirmed the osteogenic transdifferentiation of VSMCs in calcified vessels. This was evidenced by the molecular signature: decreased expression of the contractile marker α-SMA, alongside increased expression of the osteogenic regulators Runx2 and OPN (Figure 2I). This Runx2⁺/OPN⁺/α-SMA⁻ profile directly demonstrates that VSMCs switched from a contractile to a bone-forming phenotype, which is consistent with the observed histological progression from chondrogenic differentiation to overt calcification (Figure 2B-2H).

Figure 2 The cellular remodeling during the MAC progression. The CCAs of mice from the sham operation and experimental groups were collected at different time points and examined through H&E staining. Representative pictures were selected to illustrate the differentiation process of VSMC (A-H). (A) The VSMCs from the sham operation group with spindle shape were uniformly distributed in the media layer of the CCA (→). (B) Following endothelial injury to the CCA, the vessels in the experimental group exhibited intimal and medial thickening. Within these thickened areas, VSMCs proliferated, migrated, and subsequently exhibited a chondroid phenotype, identified on H&E staining by their rounded morphology, location within lacunae, and association with a basophilic extracellular matrix (→). (C) Naïve chondrocytes embarked on a proliferation program (→). (D) The formation of the initial ossification center: the naïve chondrocytes began to mature, the basophilia of the cartilage matrix grew, and a calcium salt deposition appeared, showing a bluish violet color in H&E staining (→). (E) The number of mature cartilage cells increased, and the H&E staining result showed an obvious deposition of calcium salts (→). (F) Part of the mature chondrocytes proliferated and clustered with a significant increase in size and an enhancement of the basophilia; isogenous groups were observed. (G) Hypertrophic chondrocytes began to degenerate. Blood vessels and osteoblasts invade (∆) to form osteoid, which subsequently undergoes calcification, accelerating the process of bone formation (→). (H) Bone formed inside the vascular wall, and trabecular bone was observed in the bone area, of which the surface was occupied with single osteocytes (→). (I) IHC staining of the expressions of α-SMA, OPN, and Runx2 in blood vessels of the sham operation group and experimental groups. Representative image taken at 10× objective magnification (scale bar: 200 mm). A higher-magnification view of the boxed region in, captured at 40× objective magnification. α-SMA, alpha smooth muscle actin; CCA, common carotid artery; H&E, hematoxylin and eosin; IHC, immunohistochemistry; MAC, medial arterial calcification; OPN, osteopontin; Runx2, runt-related transcription factor 2; VSMC, vascular smooth muscle cell.

IF analysis of the CCA from the MAC model group revealed substantial macrophage infiltration, characterized by significantly increased expression of both CD68 (a pan-macrophage marker) and CD86 (an M1 phenotype marker) compared to control groups (Figure 3A). This finding confirms the infiltration and polarization of macrophages toward the pro-inflammatory M1 phenotype, supporting their potential causative role in vascular calcification progression. Notably, IHC staining on adjacent sections demonstrated pronounced expression of the pro-inflammatory cytokines IL-1β and IL-6 within the calcified arterial areas. The spatial distribution of these cytokines strongly correlated with regions of macrophage accumulation identified by IF (Figure 3B). This close spatial association suggests that infiltrated M1 macrophages are likely the source of these inflammatory mediators, thereby establishing a pro-calcific microenvironment. Consequently, the resultant inflammatory milieu may promote the osteogenic transdifferentiation of VSMCs, accelerating the development of MAC.

Figure 3 Pro-inflammatory milieu in calcified arteries and the impact of guide wires with different diameters on MAC model generation. (A) Representative IF staining results showed that CD68 (green color) and CD86 (red color) expression was low in the mice from the sham operation group and significantly higher in the mice from the experimental group in terms of the fluorescence density and intensity. Individual channels are presented separately for clarity. Nuclei were counterstained with DAPI (not shown). (B) Representative IHC staining results showed that the IL-1β and IL-6 expression in the calcified blood vessel from the experimental group was significantly higher than that in the sham operation group, with slightly higher intensity. (C) Incidence rates of ossification caused by the 0.35, 0.40, and 0.45 mm diameter guide wires. (D) According to the MAC histopathological grading criteria (Table 1), the MAC grading ratios for the MAC caused by the 0.35, 0.40, and 0.45 mm diameter guide wires in the mouse MAC. Sample sizes were: n=10 for Sham group, n=12 for 0.35 mm group, and n=13 for 0.40 mm group, n=15 for 0.45 mm group. (E) Image of rough guide wires with diameters of 0.35, 0.40, or 0.45 mm. DAPI, 4',6-diamidino-2-phenylindole; IF, immunofluorescent; IHC, immunohistochemistry; IL-1β, Interleukin-1 beta; IL-6, Interleukin-6; MAC, medial arterial calcification.

Having characterized the inflammatory basis of MAC, we next aimed to establish a reproducible model by defining the key parameters for its reproducible induction. We identified guide wire size as a critical determinant, which significantly modulated calcification severity (Figure 3C-3E). Using this optimized model, we subsequently established a histopathological grading system to categorize the severity. Thus, the results below systematically outline the effects of critical parameters (guide wire size, model duration, and VD₃ supplementation) and validate the model’s responsiveness using known calcification inhibitors.

MAC pathological classification

Vascular calcification can manifest through calcium deposition, chondrogenesis, and bone formation. These pathological features may occur individually or coexist in MAC.

Since the pathological features observed in our established MAC model mirrored those seen in clinical cases, we developed a parallel, four-level grading system to categorize the severity of carotid artery calcification in mice. This grading was primarily based on the extent of calcium deposition, analogous to approaches used in clinical assessments (16-18). Calcification was identified and evaluated using histological staining, with Von Kossa staining providing specific detection of calcium phosphate deposits and H&E staining offering complementary morphological information. The classification for grades I-IV MAC was defined as the maximum diameter of calcium particle aggregation being between >0.5 and 40 µm for grade I, up to 100 µm for grade II, up to 300 µm for grade III, and ≥300 µm for grade IV (Figure 4 and Table 1).

Figure 4 The representative histopathological grading of MAC in mice. Histopathological sections from the experimental group, stained with Von Kossa (A-C) and H&E (D-J), demonstrate the progression of medial calcification severity. The corresponding grading criteria are summarized in Table 1. Insets in (A-D), (I), and (J) show higher-magnification views of the boxed areas (scale bars: 50 µm and 200 µm for main images and insets, respectively). (A) Microcalcification in Von Kossa staining under 40× magnification. (B) Punctate calcification. (C) A brown 65 μm length fragment calcification. (D) A 90 μm length fragment calcification (bluish violet) is visible with H&E staining. (E-H) represent sheet calcification, with lesion lengths measuring approximately 198 µm, 287 µm, extending to span nearly half, and nearly the entire circumference of the CCA, respectively. (I) Cartilage formation and calcium were observed: a large number of chondrocytes occupied the media in a regular arrangement; the mature chondrocytes in the central region formed isogenous groups and the slightly naïve chondrocytes were distributed individually in the periphery. (J) Bone formation and calcium deposition were observed: bone trabeculae appeared in the media, and bone cells and bone lacuna with bone marrow were visible on the surface of bone tissue. CCA, common carotid artery; H&E, hematoxylin and eosin; MAC, medial arterial calcification.

In this MAC model, calcification across grades I-IV was observed either in isolation or in conjunction with chondrogenesis and/or ossification; however, the formation of calcified nodules, as seen in the human condition, was not typically observed.

The effect of wire diameter on MAC formation

Three diameters of stainless steel rough guide wires (0.35, 0.40, and 0.45 mm) were used to scrape the vascular endothelium in mice to examine the effect of wire diameter on MAC formation in mice. A sham operation group was also established as a negative control (Figure 1A). H&E and Von Kossa staining indicated that injury to endothelial cells was rare and minor, with no calcium deposition in the blood vessels of the sham operation group. In mice, the 0.35 mm diameter stainless steel rough guide wires inflicted moderate injury to the vascular intima, while the 0.40/0.45 mm diameter stainless steel rough guide wires inflicted severe injury. The incidence rates of MAC with the 0.35, 0.40, and 0.45 mm diameter wires were 25%, 46.15%, and 86.67%, respectively. Among the cases of MAC, 8.33% and 16.67% of cases caused by the 0.35 mm diameter wires were grade I-II MAC level; 7.69%, 15.38%, 15.38%, and 7.69% of cases caused by the 0.40 mm diameter wires were grade I-V MAC; and 6.67%, 20%, 33.33%, and 26.67% of cases caused by the 0.45 mm diameter wires were grade I-V MAC. The ossification rate of the experimental group when subjected to the 0.40 and 0.45 mm diameter wires was 15.38% and 33.33%, respectively (Figure 3C-3E). Since the 0.45 mm guidewire yielded the highest incidence of MAC—which was the primary indicator of successful model establishment—it was selected as the optimal parameter for further optimization.

The impact of modeling time on MAC formation

In order to observe the MAC progression and thus determine the optimal modeling time, samples of the LCA were obtained for H&E and Von Kossa staining once a month for 4 consecutive months. In the first month, slightly thickened intima and media, damaged endothelial cells, and differentiation of smooth muscle cells were observed in the CCA in the experimental group (Figure 5A-5E). Among the experimental cases, 18.18% had MAC (grade I: 9.09%; grade II: 9.09%). In the second month, thickened intima and media in the CCA were observed in all experimental cases, and the incidence of MAC increased to 54.54% (grade I: 18.18%; grade II: 27.27%; grade III: 9.09%). Cells with morphological features of chondroid cells were observed in some experimental cases, and 18.18% of CCAs in the experimental group had ossification. In the fourth month, the MAC incidence rate (total: 81.81%; grade I: 9.09%; grade II: 9.09%; grade III: 27.27%; grade IV: 36.36%), grade IV proportion, and ossification rate (36.36%) significantly increased and were similar to those in the third month. Thus, the incidence rate of MAC appeared to be the highest at the third month, and so we selected 3 months as the modeling time in creating the MAC model in the follow-up experiments.

Figure 5 The impact of modeling time, supplementary VD₃ diet, and calcification inhibitors on the occurrence of MAC model generation. (A) The MAC incidence rate and ossification rate of the MAC model preparation with modeling times of 1-, 2-, 3-, and 4-month. (B) The MAC grading ratios of the three different experimental groups. The 0.45 mm guidewire diameter parameter set in Figure 1D is identical to that of the 3-month modeling group in this cohort and is consequently omitted for clarity. Sample sizes were: n=11 for 1-month group, n=11 for the 2-month group, and n=11 for the 4-month group. (C) Von Kossa staining showed punctate calcification in the CCA of mice from the experimental group with 1-month modeling time (→). (D) Grade III sheet calcification (191 μm) observed in a CCA from the experimental group with a 2-month modeling time (H&E staining). (E) Grade IV sheet calcification (>300 μm) observed in the vascular wall of mice from the experimental group with 3-month modeling time (H&E staining result). (F) Mechanical injury surgery and a VD₃ diet were employed to establish a mouse MAC model, and the MAC incidence rate and ossification rate were observed after the experimental mice were treated with etidronate or SNF472. (G) The MAC grading ratios of etidronate- and SNF472-treated groups. (H) After supplementation with VD₃, the proportion of calcification deposition and cartilage formation increased (H&E staining). The mature chondrocytes occupied the entire media and appeared stained as bluish violet in H&E staining, due to the cartilage matrix being strongly alkaline. Sample sizes were: n=12 for VD₃ group, n=11 for etidronate-treated group, and n=10 for SNF472-treated group. (I) Black sheet plaques almost entirely covered the media of the CCA from the VD₃ (Von Kossa staining). (J) Grade III sheet calcification (195 μm) was observed in the vascular wall of mice from the etidronate-treated group (H&E staining). (K) Grade II fragment calcification (95 μm) was observed in the vascular wall of mice from the SNF472-treated group (H&E staining). Calc, calcification; CCA, common carotid artery; H&E, hematoxylin and eosin; MAC, medial arterial calcification; VD₃, vitamin D₃.

The impact of supplementary VD₃ diet on MAC formation

In the field of vascular calcification studies, researchers often use VD₃ to induce vascular calcification (19). Therefore, we administered the mice with high-dose VD₃ continuously for 3 consecutive days after vascular injury surgery in the experimental group. The results indicated that the incidence rate of vascular MAC in mice reached 100% (grade I: 8.33%; grade II: 8.33%; grade III: 50%; grade IV: 33.34%), and the ossification rate of the MAC cases increased to 50% when the VD₃ diet was included (Figure 5F-5I). These observations indicate that VD₃ may increase both the incidence rate and severity of MAC in our mouse model. In summary, vascular injury combined with a VD₃ diet can effectively induce the calcification of the CCA in 2 months male of C57BL/6J mice.

The impact of calcium regulators on MAC formation

We investigated the effects of etidronate and SNF472 on carotid artery calcification in our MAC model. Experimental results showed that the vascular endothelial cells of mice in the etidronate and SNF472 treatment groups also sustained damage, and the intima/media experienced pathological thickening as well. However, compared with the VD₃ group, the incidence of MAC was significantly lower in the etidronate group (54.55%; grade I: 9.09%; grade II: 18.18%; grade III: 18.18%; grade IV: 9.09%; P<0.01) and the SNF472 group (50.00% reduction; grade I: 10%; grade II: 20%; grade III: 20%; grade IV: 0%; P<0.01), as determined by Fisher’s exact test. The incidence of ossification was reduced to 27.27% and 30.00%, respectively, which corresponds to inhibition rates of 45.45% and 50.00%. Furthermore, the incidence of grade IV calcification was significantly lowered to 9.09% and 0%, respectively (Figure 5F-5K). These data suggest that etidronate and SNF472 can inhibit the incidence and severity of MAC in mice of this model, and that the mouse MAC model constructed in this study may be valuable for screening calcification-inhibiting drugs.

Discussion

A vital principle in establishing an animal model is its pathogenic analogy to the human condition (20,21). In this context, we have successfully developed a novel and reproducible model of MAC in wild-type C57BL/6J mice. Distinct from models reliant on CKD or dietary perturbations, our approach induces calcification through localized and controlled mechanisms, thereby avoiding the confounding effects of systemic metabolic dysregulation. This model recapitulates key histological features of human MAC and has been functionally validated by its response to calcification inhibitors such as SNF472 and etidronate. It thus provides a well-defined and highly reproducible platform that fulfills this principle for elucidating the pathophysiology of MAC.

Using this platform, we investigated key contributors to factors of MAC formation, focusing on the roles of mechanical injury and vascular inflammation, which are important promoters of MAC formation (11,22).

To establish a robust model of carotid MAC, we applied mechanical injury to create a localized susceptible niche. The 100% success rate and significantly enhanced calcification severity (with 83.3% reaching stage II/IV and 50% exhibiting ossification) in the VD₃-combined model, compared to the 86.7% success rate with mechanical injury alone, strongly suggests that the two stimuli act on complementary pathways. This combinatorial strategy was designed to overcome the limitations of VD₃-alone models, which predominantly induce calcification in the elastic aorta but show limited efficacy in the CCA (13,23). These findings strongly suggest synergistic and complementary interactions between the two stimuli.

Mechanical injury initiates the process by inducing endothelial disruption and cytoskeletal disassembly in VSMCs, thereby activating pro-inflammatory mechanosensitive pathways such as integrin-mediated signaling (24). Cellular studies indicate that mechanical stress disrupts the cytoskeleton and mechanosensitive pathways in VSMCs, involving integrins, actin, and calponin. This disruption promotes a phenotypic switch toward a synthetic state, characterized by enhanced proliferation and migration, along with reduced α-SMA expression. Through mechanosensing and mechanotransduction, VSMCs maintain and remodel the extracellular matrix (25,26). However, under pathological conditions, this process leads to chondrocyte-like phenotypic transition and ultimately facilitates vascular calcification.

VD₃ does not simply accelerate this process but acts synergistically by exacerbating endothelial dysfunction and amplifying the inflammatory response. Furthermore, VD₃ directly promotes osteogenic transdifferentiation of VSMCs, as supported by the upregulation of key markers such as Runx2 in VD₃-induced models. This synergy likely involves multiple mechanisms: VD₃ elevates the serum calcium-phosphate product, facilitating hydroxyapatite deposition, while concurrently activating intracellular signaling cascades that converge with pathways triggered by mechanical injury (27,28). Together, these synergistic insults robustly initiate and amplify a pathological program of inflammation and osteogenic differentiation, leading to the highly efficient and consistent calcification observed in our model.

In our mouse model of MAC, we observed a similar process of osteogenic calcification (Figure 2A-2H). Consistent with this finding, MAC-affected mice showed elevated expression of the bone formation-related proteins OPN and Runx2 (Figure 1I). Clinically, grade II-IV vascular calcification is often accompanied by ossification (16). Similarly, in our MAC model, higher-grade calcified vessels frequently exhibited ossification, indicating that the severity of calcification is associated with the likelihood of osteogenic transition. These findings align with the understanding that chronic low-grade inflammation represents a critical stage in the progression of vascular calcification (29).

Concurrently, macrophages exacerbate arterial calcification by phagocytosing calcific debris and releasing proinflammatory cytokines such as IL-1β and IL-18. This response perpetuates vascular injury and calcification, while chronic inflammation induces oxidative stress, further damaging the endothelium (30,31). In our model, macrophages infiltrating the damaged vascular wall exhibited a pro-inflammatory phenotype, marked by elevated release of M1-associated cytokines, including IL-1β and IL-6 (Figure 3A,3B). This suggests a polarization toward an M1-like state, potentially aggravating vascular injury. It should be noted, however, that definitive characterization of macrophage polarization will require future analysis of specific surface markers [e.g., inducible nitric oxide synthase (iNOS), cluster of differentiation 206 (CD206), arginase-1]. These results suggest that our model can effectively simulate the pathogenic process of clinical vascular calcification.

While our acute injury model does not recapitulate the chronic progression of human MAC, it provides a powerful and reproducible system to delineate its initiating mechanisms. We posit that core drivers of calcification—such as inflammatory cell recruitment, VSMC phenotypic switching, and hydroxyapatite nucleation—are conserved across acute and chronic contexts (32). This model is uniquely suited to resolve these early events, which are difficult to capture in slower, chronic models. Future studies will leverage this platform to identify key initiating factors for subsequent validation in chronic models of aging or CKD.

Our study has limitations that should be acknowledged. First, the high-dose VD₃ required for robust calcification may cause systemic ectopic calcification, which was not evaluated, and the optimal dosage remains undefined. Future studies need to titrate VD₃ and assess off-target effects. Second, the lack of quantitative measurements of medial and intimal thickness limits insight into vascular remodeling; future morphometric analyses will address this. Notwithstanding this utility, we caution against the direct translation of our findings to human disease. Species-specific differences in vascular architecture, hemodynamics, and repair mechanisms preclude direct extrapolation. Thus, while our model offers an invaluable controlled platform for mechanistic discovery, validating the translational relevance of the identified pathways will require future studies in more complex systems, including those utilizing human tissues or chronic disease settings.

Conclusions

We established an efficient mouse MAC C57BL/6J mouse model. The pathological process and symptoms of this MAC model are similar to those of clinical practice. Additionally, we developed a mouse MAC histopathological grading system based on this MAC model, providing a semiquantitative standard for the potential application of the model in the future. This model is expected to have broad application prospects in both basic medicine and drug screening research.

Acknowledgments

We appreciate those who were involved in this study. We would like to thank Sihong Yu for her help in editing the language of the manuscript.

Footnote

Reporting Checklist: The authors have completed the ARRIVE reporting checklist. Available at https://cdt.amegroups.com/article/view/10.21037/cdt-2025-435/rc

Data Sharing Statement: Available at https://cdt.amegroups.com/article/view/10.21037/cdt-2025-435/dss

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Funding: This research was funded by the Natural Science Foundation of Zhejiang Province, China (No. LGF22H300007).

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://cdt.amegroups.com/article/view/10.21037/cdt-2025-435/coif). M.Z. is an employee of Huzhou Talewell Biopharmaceutical Co., Ltd. Huzhou, China. The other authors have no conflicts of interest to declare.

Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. All animal experiments were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. This study was approved by the Animal Welfare and Ethics Committee of Huzhou University (No. 20220701).

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

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