A murine model of aortic regurgitation generated by trans‐apical wire destruction of the aortic valve

Abstract

Background

The mechanisms underlying cardiac remodeling in aortic valvular (AoV) disease remain poorly understood, partially due to the insufficiency of appropriate preclinical animal models. Here, we present a novel murine model of aortic regurgitation (AR) generated by trans‐apical wire destruction of the AoV.

Methods

Directed by echocardiography, apical puncture of the left ventricle (LV) was performed in adult male C57BL/6 mice, and a metal guidewire was used to induce AoV destruction. Echocardiography, invasive LV hemodynamic and histological examination were conducted to assess the degree of AR, LV function and remodeling.

Results

AR mice exhibited rapid aortic regurgitation velocity (424 ± 15.22 mm/s) immediately following successful surgery. Four weeks post‐surgery, echocardiography revealed a 54.6% increase in LV diastolic diameter and a 55.1% decrease in LV ejection fraction in AR mice compared to sham mice. Pressure‐volume catheterization indicated that AR mice had significantly larger LV end‐diastolic volumes (66.2 ± 1.5 μL vs. 41.8 ± 3.4 μL), reduced LV contractility (lower dP/dt_max and Ees), and diminished LV compliance (smaller dP/dt_min and longer Tau) compared to sham mice. Histological examination demonstrated that AR mice had significantly larger cardiomyocyte area and more myocardial fibrosis in LV tissue, as well as a 107% and a 122% increase of heart weight/tibial length and lung weight/tibial length, respectively, relative to sham mice.

Conclusions

The trans‐apex wire‐induced destruction of the AoV establishes a novel and efficient murine model to develop AR, characterized by significant eccentric LV hypertrophy, heart failure, and pulmonary congestion.

Directed by echocardiography, apical puncture of the left ventricle (LV) was performed in adult male C57BL/6 mice, and a metal guidewire was used to destroy the aortic valve. This model can cause larger scale regurgitation, significant eccentric LV hypertrophy, heart failure and pulmonary congestion.

1. INTRODUCTION

Aortic regurgitation (AR) is a common form of valvular heart disease that causes left ventricular volume overload and, ultimately, heart failure. Surgical or transcatheter aortic valve (AoV) replacement is the only current treatment for AR‐induced heart disease, while existing pharmacologic therapy alone cannot improve the prognosis of severe AR. ¹ , ² The mechanisms underlying cardiac remodeling in AR and the reversal of remodeling following AoV replacement remain largely unclear, partly due to the lack of appropriate preclinical animal models. Murine models of AR are scarce, creating a translational knowledge gap between understanding the mechanisms and developing new pharmacologic therapies.

Two types of murine AR models have been reported: (1) gene‐modified mice, such as natriuretic peptide receptor 2/low‐density lipoprotein receptor deficient mice, ³ hypomorphic epidermal growth factor receptor mutation mice (Wave mice) ⁴ and Naïve New Zealand obese mice ⁵ ; and (2) mice with surgically induced AoV destruction. However, gene‐modified models often exhibit a low incidence of heart failure phenotype or require a prolonged period to develop heart failure. Additionally, a previous report showed that a surgical AR mouse model generated by trans‐carotid artery approach induced only moderate volume overload and heart failure without significant pulmonary congestion (a 20% increase in left ventricular (LV) diameter and a 30% decrease in LV fractional shortening (LVFS) at 4 weeks after surgery). ⁶

Given the limitations of the trans‐carotid AR model, including the loss of a carotid artery and insufficient volume overload, there is room for modification. In this study, we introduce a trans‐apical AR murine model characterized by substantial aortic reflux, significant LV volume overload, markedly depressed systolic and diastolic function of the LV, and pronounced pulmonary congestion.

2. METHODS

2.1. Generation of AR through apex‐LV‐AoV approach

All procedures were performed in accordance with our institution's guidelines for animal research that conform to the Guide for the Care and Use of Laboratory Animals (National Institutes of Health Publication, 8th Edition, 2011). This study was approved by the ethics committee of Nanfang Hospital, Southern Medical University.

C57BL/6 male mice (8–10 weeks old and weighing 20–25 g) were anesthetized by intraperitoneal injection of a mixture of xylazine (5 mg/kg) and ketamine (100 mg/kg), and their breathing was supported by a ventilator. A left‐sided thoracotomy was then performed between the fourth and fifth ribs to expose the heart (echocardiography‐directed percutaneous puncture of LV without chest opening was also feasible). A 26‐gauge needle was used to puncture the apex of the LV, and then the needle was withdrawn after entering the LV chamber. Guided by echocardiography, a metal PTCA (percutaneous transcoronary angioplasty) guidewire (diameter: 0.36 mm, ASAHI, Japan) was introduced through the apical puncture site into the LV chamber (Figure 1A). The guidewire was then manipulated to advance into the LV outflow tract and then repeatedly puncture the AoV 1–3 times at its early opening stage (early systolic period) to damage the AoV (Figure 1B), until significant diastolic regurgitation was confirmed by Doppler echocardiography in the thoracic aorta or under the AoV. Damaging the AoV is a critical step in this model: the valvular puncture timing should not be during the diastolic period (overcoming the great resistance to puncturing would lead to large AR and acute pulmonary edema), nor during the late systolic period (as the guidewire could easily to pass through the AoV without causing damage). Finally, the guidewire was removed, and the chest was closed with 5–0 silk sutures. Sham mice underwent the same procedures, except for the AoV damage step.

FIGURE 1.

Schematic of apical approach for generation of mouse AR model. (A) Surgical schematic and procedure: 1. skin cutting in apical zone; 2. exposure of fourth and fifth ribs; 3. exposure of heart apex; 4. a 26‐gauge needle was used to puncture the apex; 5. a metal guidewire was introduced into LV chamber through the apical puncture site. (B) Echocardiography‐guided wire to injure the aortic valve: 1. long‐axis imaging of LV showing clearly the LV chamber and aortic valve (red arrow); 2. visible guidewire in LV chamber (blue arrow); 3. guidewire reaching aortic valve. AR, aortic regurgitation; LV, left ventricle.

2.2. Echocardiography

Echocardiography was performed during the operation and post‐operation using a Vevo 2100 system (Fujifilm Visual Sonics, Ontario, Canada). As previously described, ⁷ , ⁸ mice were anesthetized with isoflurane (3% for induction, 1.5% for maintenance), and the heart rate was maintained between 450 and 550 beats/min. Long‐axis images of the LV were obtained under B‐mode, and short axis views at the papillary muscle level were recorded under M‐mode. LV diastolic and systolic posterior wall thickness (LVPWd, LVPWs), LV end‐diastolic and systolic diameters (LVEDd, LVESd), and LV ejection fraction and fractional shortening (LVEF, LVFS) were measured by adjusting the measurement interface to assess cardiac function. Color Doppler mode was used to measure the peak systolic and diastolic flow velocity of thoracic aorta (PSFV and PDFV).

2.3. Hemodynamic examination

The LV pressure‐volume (PV) data were collected 4 weeks after AR surgery. Conductance and pressure signals were obtained using a SPR‐839 1.4F Millar PV catheter (Millar Instruments, Inc., Houston, TX, USA). Following a previously described protocol, ⁹ mice were anesthetized with ketamine/xylazine (light anesthesia with 80/10 mg/kg), and the PV catheter was inserted into the ascending aorta and LV via the right common carotid artery. Heart rate (HR), LV systolic and end‐diastolic pressure (LVSP and LVEDP), maximum derivative of LV pressure and volume at systolic and diastolic period (±LV dP/dt_max, ±LV dV/dt_max), end‐diastolic volume (EDV), stroke volume (SV) and cardiac output (CO), stroke work (SW), exponential time constant of relaxation (Tau), end‐systolic elastance (Ees), and slope of end‐diastolic pressure volume relationship (EDPVR) were analyzed with PowerLab software (PV loop module; AD Instruments, Australia).

2.4. Histological examination

Mice were euthanized 4 weeks after AR or sham surgery. The hearts were harvested, fixed in 4% paraformaldehyde, and paraffin‐embedded according to standard protocols. Hematoxylin–eosin (H&E) and wheat germ agglutinin (WGA) staining were used to evaluate aortic valve morphology and myocardial hypertrophy, respectively. Masson's trichrome stain was used to assess myocardial fibrosis.

2.5. Statistical analysis

All statistical analyses were performed using GraphPad Prism 9.0 (GraphPad Software, San Diego, CA, USA) and summarized by presenting mean ± standard error of the mean (SEM). Statistical differences between two groups were evaluated using two‐tailed, unpaired t‐tests. Survival rates were calculated according to the Kaplan–Meier method. P values < 0.05 were considered statistically significant.

3. RESULTS

3.1. Trans‐apical approach induces large scale reflux

The extent of AoV damage was judged by the severity of AR detected with Doppler ultrasound. In AR mice, color Doppler imaging at the aortic arch revealed a significant large‐scale reflux (PDFV: 424.1 ± 15.2 mm/s), which was absent in sham mice. Additionally, the forward flow velocity at systole in AR mice was 37.9% higher than in sham mice (PSFV: 1021.9 ± 26.2 mm/s vs. 741.0 ± 16.9 mm/s) (Figure 2A–C). Similar results were obtained when the probe was positioned on the LV (Figure 2D). The perioperative mortality was 20%, and the survival rate was 75% at 4 weeks in mice with successful AR surgery (Figure 2E).

FIGURE 2.

Echocardiographic confirmation of the success of AR generation. (A) Doppler images for measurement flow velocity at the aortic site. In the 2D images of aortic arch at diastolic period, a stream of colorful reflux was detected in AR mice. (B) The peak systolic flow velocity in thoracic aorta (PSFV). (C) The peak diastolic velocity in thoracic aortic (PDFV). (D) Doppler images of a reflux entering from ascending aorta (AO) into LV at long axis view. (E) Survival rate of AR mice at 4 weeks after surgery. Data are means ± SEM. *P < 0.05 vs. sham group. AR, aortic regurgitation; LV, left ventricle.

3.2. Cardiac remodeling evaluated by echocardiography

At 4 weeks after surgery, echocardiography demonstrated that AR mice had significantly larger diastolic LV wall thickness (LVPWd and LVAWd) and LV diameters (LVEDd and LVESd) (Figure 3A–E), and significantly reduced LVEF (55.1% lower) and LVFS (61.5% lower) compared to the sham group (Figure 3F,G). Estimated diastolic LV volume in the AR group was 173.5% larger than in the sham group (183.1 ± 6.4 μL vs. 66.9 ± 3.3 μL) (Figure 3H), while LVPWd was significantly smaller in the AR group than in the sham group. These findings indicate the presence of significant eccentric hypertrophy in AR mice.

FIGURE 3.

Echocardiographic results at 4 weeks after AR surgery. (A) Representative M‐mode echocardiography of LV. (B) LV diastolic posterior wall thickness (LVPWd). (C) LV systolic posterior wall thickness (LVPWs). (D) LV end‐diastolic diameter (LVEDd). (E) LV end‐systolic diameter (LVESd). (F) LV ejection fraction (LVEF). (G) LV fractional shortening (LVFS). (H) Calculated LV volume. Data are means ± SEM. *P < 0.05 vs. sham group. AR, aortic regurgitation; LV, left ventricular (or ventricle).

3.3. Hemodynamic changes in AR mice

PV catheter examination indicated similar LVSP between AR mice and sham mice (Figure 4A,B). Compared to the sham group, AR mice exhibited similar LVEDP and HR (Figure 4C,D), higher SV and CO (Figure 4E,F), increased cardiac workload (SW, increased by 128%) (Figure 4G), greater volume load (LVEDV and LV change rate dV/dt_max were increased by 58.6% and 141.7%, respectively) (Figure 4H,I). Additionally, AR mice exhibited impaired systolic function, with a decrease in LV dP/dt_max and Ees by 37.2% and 49.2%, respectively. Diastolic function was also compromised, as evidenced by a 42% reduction in LV dP/dt_min, and an increase in the EDPVR slope and Tau (97% and 88.9% higher, respectively) (Figure 4J–N). These findings suggest that trans‐apical AR‐induced LV volume overload in mice leads to both systolic and diastolic dysfunction, as well as a marked increase in LV work.

FIGURE 4.

LV hemodynamic alterations at 4 weeks after AR surgery. (A) Representative LV pressure–volume loops during vena cava occlusion; (B) LV systolic pressure (LVSP). (C) Heart rate (HR). (D) LV end‐diastolic pressure (LVEDP). (E) Stroke volume (SV). (F) Cardiac output (CO); (G) Stroke work (SW). (H) End‐diastolic volume (EDV). (I) Maximum rate of volume increase in LV (dV/dt max). (J) Maximum rising rate of LV pressure (dP/dt_max). (K) LV end‐systolic elastance (Ees). (L) Maximum descending rate of LV pressure (dP/dt_min). (M) Exponential LV relaxation time constant (Tau). (N) End‐diastolic pressure‐volume relationship as a measure of LV diastolic stiffness (inverse of compliance) (slope of EDPVR). Data are means ± SEM. *P < 0.05 vs. sham group. AR, aortic regurgitation; LV, left ventricular (or ventricle).

3.4. AR induces eccentric hypertrophy and pulmonary congestion

Hearts and lungs of mice were harvested 4 weeks after surgery. AoV damage was verified by histological examination, with thrombus formation visible in the damaged AoV (Figure 5A). Body weight was similar between AR and sham mice (Figure 5C), while morphological examination showed AR mice had markedly larger hearts and dilated LV compared to sham mice (Figure 5B). The heart weight to body weight (HW:BW) and HW to tibial length (HW:TL) ratios in AR mice were 107% and 113% greater, respectively, than sham mice (Figure 5D,E). Larger cardiomyocyte area and more myocardial fibrosis were found in AR mice (Figure 5F–H). Moreover, severe pulmonary congestion characterized by a significant increase in lung weight to body weight (LW:BW) and LW to tibial length (LW:TL) ratios (130% and 122% respectively larger than sham mice) was confirmed in AR mice (Figure 5I–K).

FIGURE 5.

Histological changes at 4 weeks after AR surgery. (A) Representative H&E staining images of mouse aortic valves. (B) Representative whole mouse hearts and HE staining of heart cross‐sections. (C) Body weight (BW). (D) HW:BW ratio; (E) HW:TL ratio. (F) Representative myocardial cells stained with WGA and myocardial cross‐sections stained with Masson's trichrome. (G) Quantization of cardiomyocyte cross‐sectional area. (H) Quantization of myocardial fibrosis. (I) Representative whole mouse lungs. (J) LW:BW ratio. (K) LW:TL ratio. Data are means ± SEM. *P < 0.05 vs. sham group. AR, aortic regurgitation; LV, left ventricular (or ventricle); HW, heart weight (mg); LW, lung weight (mg); TL, tibial length (mm); HE, hematoxylin–eosin; WGA, wheat germ agglutinin.

4. DISCUSSION

This is the first report to describe a novel approach for constructing a murine AR model. Unlike the forward approach (trans‐carotid artery), damaging the AoV by trans‐apical wire puncture (reverse approach) induces more extensive regurgitation, greater LV volume overload, more severe heart failure, and significant pulmonary congestion. Moreover, this method avoids the loss of a carotid artery, which could lead to brain hypoperfusion. The trans‐carotid AR mouse model induces approximately a 20% increase in LV diameter and a 30% decrease in LVFS. ⁶ In contrast, our trans‐apical approach‐induced AR resulted in a 54% increase in LV diameter and a 61% decrease in LVFS. The survival rate between the two approaches was similar.

AR is a common condition in clinical practice, often resulting from age‐related degenerative changes, rheumatic heart disease, or severe LV dilation. Persistent AR leads to progressive cardiac remodeling and heart failure. Surgical or transcatheter AoV replacement is the mainstay of treatment for AR. However, cardiac remodeling cannot be efficiently reversed by AoV replacement in some patients, highlighting the unmet clinical need for developing new pharmacotherapeutic strategies to prevent and reverse AR‐induced cardiac remodeling. Ideal AR models should closely mimic the clinical scenario, facilitating a better mechanistic understanding and the testing of novel treatment strategies for AR disease.

Several rodent AR models have been reported. In rats with surgically induced AR, a significant reduction of LVFS takes approximately 12 weeks to develop. ¹⁰ Naïve New Zealand obese (NZO) mice have the possibility of spontaneously developing degenerative AoV stenosis and regurgitation, and need about 22 weeks to develop end‐stage heart failure. ⁵ Genetic models, such as Wave mice and Krox20 (Egr‐2) deletion mice, can induce AoV hypoplasia or dysfunction. ⁴ , ¹¹ Additionally, the high‐fat diet‐induced AR mouse model typically requires 6–9 months to develop the AR phenotype, with an approximate 75% success rate. ¹²

The aortocaval shunt model is the most commonly used volume overload model, which increases cardiac preload by anastomosing the abdominal aorta and inferior vena cava. ¹³ However, this model does not accurately simulate the clinical condition. The surgical AR mouse model, created by metal guidewire puncture of the AoV is economical and practical. AoV injury, induced by inserting a guidewire into the mouse LV via the right common carotid artery (RCCA), can result in AoV stenosis and/or AR. ⁶ , ¹⁴ , ¹⁵ However, our experience indicates that this approach does not achieve a high incidence of severe heart failure due to insufficient volume overload. In addition, this surgical procedure requires ligation of the RCCA, which can negatively affect cerebral blood supply. Previous studies have confirmed that unilateral carotid artery ligation induces chronic cerebral hypoperfusion in mice, ¹⁶ and this procedure is also a well‐established model for cerebral ischemic injury in immature mice. ¹⁷ Increasing evidence links brain damage with cardiac dysfunction. ¹⁸ In contrast, the apical approach overcomes these disadvantages, and reduces operation time by omitting arterial cannulation. A key aspect of this technique is the timing of AoV puncture: the guidewire puncture should be performed at the initial stage of AoV opening (early systolic period) as guided by long‐axis echocardiographic images of the LV. Puncturing during the diastolic period would require overcoming substantial resistance, potentially leading to excessively large AR and acute heart failure. Although the apex puncture may cause slight bleeding, the wound is usually sealed quickly due to myocardial contraction.

The progression of cardiac remodeling in patients with AR is a chronic process, with the severity of AR typically increasing over time. In contrast, the degree of AR induced mechanically in mice remains relatively stable. In this study, we only reported data observed 4 weeks post‐AR surgery. Fortunately, our preliminary experiments indicated that 8‐week post‐AR mice had further progression in LV remodeling (heart weight increased from about 120 mg in sham to 300 mg in AR mice), heart failure and mortality. Additionally, the perioperative mortality in our model was approximately 25%, which is lower than that observed in mouse myocardial infarction models. Moving forward, we are exploring less invasive surgical techniques. We have tried a new approach to avoid chest‐opening surgery through ultrasound‐guided percutaneous puncturing LV to damage the AoV with a guide wire.

5. CONCLUSIONS

The apical approach AR mouse model stably induces sufficient LV volume overload to develop heart failure with significant lung congestion. The severity of AR progressively increases in this model, which is more consistent with the clinical setting.

AUTHOR CONTRIBUTIONS

Xiaoxia Huang: Data curation; formal analysis. Qiancheng Wang: Data curation; formal analysis. Dan Han: Formal analysis. Hairuo Lin: Conceptualization; validation. Zhihong Li: Data curation. Cankun Zheng: Formal analysis; visualization. Jianping Bin: Validation. Wangjun Liao: Validation. Zhanchun Cong: Conceptualization; validation; writing – review and editing. Mengjia Shen: Conceptualization; validation; visualization; writing – original draft. Yulin Liao: Supervision; writing – review and editing.

FUNDING INFORMATION

This work was supported by grants from the National Natural Science Foundation of China (82370242, 82272602 and 82100407), the Natural Science Foundation of Guangdong Province (2023A1515110032 and 2022A1515220152), and Guangzhou Key Research and Development Program Foundation (202206010199).

CONFLICT OF INTEREST STATEMENT

The authors declare that they have no conflict of interest.

ETHICS STATEMENT

All procedures were performed in accordance with our institution's guidelines for animal research that conform to the Guide for the Care and Use of Laboratory Animals (National Institutes of Health Publication, 8th Edition, 2011). This study was approved by the ethics committee of Nanfang Hospital, Southern Medical University (Ethical approval number: K2021011).

Huang X, Wang Q, Han D, et al. A murine model of aortic regurgitation generated by trans‐apical wire destruction of the aortic valve. Anim Models Exp Med. 2025;8:493‐500. doi: 10.1002/ame2.12558