An Optimized Protocol for Guidewire-Induced Aortic Valve Stenosis Model in Mice

Summary

This protocol details an optimized guidewire-induced aortic valve stenosis model in C57BL/6 mice. A custom 3D-printed vascular support platform enables precise hemorrhage control, yielding 95% 24-hour survival (19/20) and 90% overall model success at 4 weeks (18/20).

Abstract

Aortic valve stenosis (AVS) is a major cardiovascular disease with increasing prevalence in aging populations, yet the scarcity of diseased valve tissue severely limits mechanistic studies and therapeutic development. This protocol establishes and optimizes a reproducible mouse model of AVS by improving upon the Honda guidewire-induced valve injury method through controlled hemorrhage management. Male and female C57BL/6 mice undergo guidewire-induced aortic valve injury; the procedure is refined by a custom 3D-printed vascular support platform that achieves precise hemorrhage control during guidewire insertion and withdrawal. Outcome measures include 24 h survival, echocardiographic assessment at 4 weeks, and histopathological validation. The optimized protocol achieves a 95% 24 h survival rate (19 of 20 animals); among survivors, 94.7% (18 of 19) meet the predefined hemodynamic success criterion at 4 weeks, yielding an overall model success rate of 90% (18 of 20 animals), confirmed by echocardiography and histological analysis. This standardized model provides a reliable, reproducible platform for investigating mechanisms of valve degeneration and evaluating candidate therapeutic interventions.

Introduction

Valvular heart disease remains a substantial healthcare burden in both developed and developing nations, with prevalence rising in parallel with population aging¹. Although transcatheter aortic valve implantation (TAVI) has transformed the treatment of severe aortic stenosis^2,3, the cellular and molecular mechanisms that drive valve degeneration are still incompletely understood, and no approved pharmacological therapy exists to slow or reverse disease progression^4,5. A central obstacle in valve disease research is the limited availability of diseased tissue for investigation^6,7. Human aortic valve specimens are difficult to obtain in sufficient quantity, and animal models that reliably recapitulate pathological valve changes have been equally scarce⁸. This gap has driven sustained efforts to develop animal models that reproduce the hemodynamic and histological features of aortic valve stenosis (AVS).

Conventional dietary approaches, such as prolonged high-fat feeding in ApoE⁻/⁻ mice, require 20–24 weeks and yield low success rates, with only a minority of animals developing a definitive stenotic phenotype^9,10,11,12. The guidewire-induced valve injury model described by Honda et al. addressed many of these shortcomings by mechanically injuring valve leaflets through carotid artery access¹³. The original technique involves carotid artery dissection, retrograde guidewire passage to the aortic valve, repeated leaflet scratching, and wire rotation—producing rapid hemodynamic changes within days and a stable stenotic phenotype by 4 weeks^13,14,15,16. Because of its relative speed and directness, the Honda method has been widely adopted in recent valve research. However, several important gaps remain. Complete procedural details have not been fully documented in the literature, standardization across laboratories is lacking, and the role of hemorrhage control—a major determinant of perioperative morbidity in this model—has never been systematically addressed. These inconsistencies reduce reproducibility and limit clinical translatability.

It was hypothesized that systematic optimization of hemorrhage control could meaningfully improve perioperative survival and model reliability. This protocol presents a refined and standardized approach that builds on the Honda method by introducing a custom-designed 3D-printed vascular support platform. The device enables adjustable longitudinal tension on the carotid artery, providing complete temporary hemostasis during guidewire insertion and withdrawal without crushing the vessel. With this approach, the optimized protocol achieves a 95% 24 h survival rate (19 of 20 animals); among survivors, 94.7% (18 of 19) meet the predefined hemodynamic success criterion at 4 weeks, yielding an overall model success rate of 90% (18 of 20 animals). Step-by-step procedural documentation is provided to facilitate reproducibility across laboratories.

Protocol

NOTE: All animal procedures described in this protocol were performed in accordance with the Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee (IACUC) of West China Hospital (Protocol No. 20240311055). Male C57BL/6 mice (8–10 weeks; approximately 22–25 g) and female C57BL/6 mice (8–10 weeks; approximately 18–20 g) were used throughout. All materials, reagents, instruments, software, and custom devices used in this protocol, including the custom 3D-printed vascular support platform, are listed in the Table of Materials.

1. Pre-operative preparation and surgical incision

1. Anesthetize the mouse using isoflurane (3% for induction, 1.5–2% for maintenance) vaporized in 100% oxygen. Confirm adequate anesthetic depth by verifying the absence of pedal withdrawal reflexes (toe pinch).
   CAUTION: Isoflurane is a volatile anesthetic. Use it with active scavenging or in a well-ventilated area, wear appropriate PPE, and follow institutional safety procedures.
2. Apply ophthalmic ointment to both eyes. Place the mouse in a supine position on a temperature-regulated surgical platform maintained at 37 °C.
3. Prepare the surgical field by depilating the anterior cervical region with chemical hair removal cream.
4. Disinfect the skin with three alternating cycles of povidone-iodine (or chlorhexidine) and 70% alcohol (or warm sterile saline), finishing with the antiseptic.
5. Infiltrate 0.5% lidocaine subcutaneously along the planned incision line for pre-emptive local analgesia.
6. Then make a longitudinal midline skin incision (approximately 1.0–1.5 cm) on the anterior neck, positioned slightly to the right of the tracheal midline, using micro-scissors.
   CAUTION: Micro-scissors, needles, guidewires, and other sharps may cause injury. Handle them carefully and dispose of used sharps in approved sharps containers.

2. Vessel isolation and suture placement

1. Under a surgical stereomicroscope, expose the right common carotid artery (RCCA) by blunt dissection through the overlying fascia and muscle layers. Carefully separate the RCCA from the adjacent vagus nerve and surrounding connective tissue, taking care to avoid mechanical irritation of the vagus nerve.
   NOTE: Keep the salivary glands and exposed tissues moist with warm sterile saline throughout the procedure to prevent desiccation.
2. Permanently ligate the distal end of the RCCA with a 6-0 silk suture tied as a secure non-slip knot.
3. Place a loose slipknot of 6-0 silk suture around the proximal segment of the RCCA, leaving it untightened for later use during guidewire insertion.

3. Platform application and arteriotomy

NOTE: Before surgery, sterilize the 3D-printed platform using ethylene oxide (EtO) gas, followed by adequate aeration, and sterilize all metal surgical instruments by autoclaving.

CAUTION: Ethylene oxide is toxic, flammable, and carcinogenic. Use EtO sterilization only in approved facilities, allow adequate aeration, and follow institutional waste-handling procedures.

1. Carefully insert the 3D-printed vascular support platform underneath the isolated segment of the RCCA, positioning the inverted V-shaped apex directly beneath the vessel.
2. Slowly adjust the platform to increase longitudinal tension on the RCCA by engaging the pre-placed sutures into progressively higher side notches.
   NOTE: The inverted V-shaped apex is designed at a 120° angle to conform to the vessel's curvature without crushing the wall. Stop increasing tension at the exact point when arterial pulsation is no longer visually perceptible. This critical visual endpoint ensures temporary hemostasis without excessive vessel injury.
3. Make a small transverse arteriotomy (approximately half the vessel circumference) on the stabilized RCCA segment using micro-scissors. Minimal or no bleeding should occur if the tension has been set correctly.

4. Guidewire insertion and valve injury

1. Under microscopic visualization, insert the guidewire (0.014 inch / 0.36 mm diameter) into the arterial lumen through the transverse arteriotomy in a retrograde direction toward the aortic root.
   NOTE: This caliber is standard to published guidewire-injury models and approximates the luminal diameter of the murine RCCA (~0.4–0.5 mm), giving a guidewire-to-vessel ratio of approximately 0.7–0.9.
   CAUTION: The guidewire can puncture vessels or surrounding tissues. Advance it slowly under visualization and dispose of used guidewires as sharps or contaminated surgical waste.
2. Tighten the pre-placed proximal slipknot around both the vessel and the guidewire to achieve complete hemostasis at the insertion site. Once the slipknot is secured, carefully remove the 3D-printed platform from beneath the vessel.
   NOTE: If unexpected bleeding occurs during insertion, or if the guidewire diameter creates excessive friction, temporarily reposition the platform to assist with hemostasis, or reduce platform tension slightly to allow the vessel lumen to expand before re-attempting insertion.
3. Advance the guidewire carefully in a retrograde direction through the RCCA, past the aortic arch, and toward the aortic valve. Under real-time echocardiographic guidance, confirm that the guidewire tip has crossed the aortic valve and entered the left ventricle.
4. With the proximal slipknot fully tightened to maintain hemostasis, perform the mechanical valve injury by executing 100 push-pull (back-and-forth) motions across the aortic valve, followed by 100 rotational motions of the guidewire against the valve leaflets.
   NOTE: The 100 push-pull and 100 rotational excursions were established through extensive in-house optimization as the numbers that best balance procedure time with reliable induction of hemodynamically significant stenosis.

5. Guidewire withdrawal and wound closure

1. Upon completion of the valve injury, slowly withdraw the guidewire from the vessel.
2. Immediately convert the proximal slipknot into a permanent non-slip knot to ligate the proximal RCCA and prevent any post-procedural bleeding.
3. Close the skin incision with interrupted 5-0 nylon sutures or surgical wound clips.
4. Do not administer post-operative non-steroidal anti-inflammatory drugs (NSAIDs). Do not administer perioperative antibiotics, as the procedure is performed under aseptic conditions.
5. Transfer the animal to a clean, warmed recovery cage and monitor continuously until full ambulation is restored.

6. Echocardiographic evaluation of transvalvular hemodynamics

1. Perform echocardiographic assessments on all surviving mice at 4 weeks post-surgery to evaluate aortic valve stenosis severity.
2. Induce anesthesia by placing the animal in an induction chamber with 3% isoflurane in 100% oxygen. Confirm anesthetic depth by verifying the loss of pedal withdrawal reflexes. Depilate the anterior chest wall using chemical hair removal cream. Apply ophthalmic ointment to both eyes to prevent corneal desiccation.
3. Transfer the anesthetized mouse to a temperature-regulated imaging platform in a supine position.
4. Secure the paws to surface electrocardiogram (ECG) electrodes for continuous heart rate monitoring. Titrate isoflurane concentration (typically 1.0–1.5%) to maintain a stable physiological heart rate of 450–550 beats per min.
5. Clean the anterior chest wall and apply a generous layer of pre-warmed acoustic coupling gel.
6. B-mode imaging acquisition
   1. Using a Vevo 3100 ultrasound system with an MX550D transducer (40 MHz), press the B-Mode control to begin live two-dimensional imaging.
   2. Position the transducer in the left parasternal position to obtain a parasternal long-axis view of the left ventricle, aortic valve, and ascending aorta. Optimize depth, width, and 2D gain.
7. M-mode acquisition and left ventricular measurement
   1. From the parasternal long-axis view, press the M-Mode control. Position the sampling line across the left ventricle at the papillary-muscle level.
   2. Press Start to begin acquisition. Once a stable trace spanning at least three cardiac cycles is obtained, press Save.
   3. In Vevo LAB, use the LV Trace tool to trace the endocardium at end-diastole and end-systole, and record the left ventricular internal dimensions, anterior and posterior wall thicknesses (LVAWd, LVPWd), and left ventricular mass.
8. Reposition and angle the transducer to acquire a dedicated aortic arch view.
   ​NOTE: Obtaining this specific aortic arch view is essential for accurate transvalvular velocity measurements, as it ensures optimal alignment of the ultrasound beam with the direction of blood flow across the aortic valve.
9. Color Doppler assessment
   1. From the aortic arch view, press the Color control. Position the color box over the aortic valve and proximal ascending aorta.
   2. Adjust the color scale and gain so the systolic jet is clearly shown without aliasing. Press Start, then Save.
10. Pulsed-wave Doppler velocity measurement
    1. Press the PW Doppler control. Position the sample-volume gate at the narrowest point of the jet (vena contracta), and apply angle correction (<60°).
    2. Press Start to begin the spectral trace; once at least three cardiac cycles are recorded, press Save. In Vevo LAB, measure peak transvalvular velocity with the Peak Velocity tool, averaging at least three cycles.

7. Histological processing and aortic root staining

1. Following echocardiographic evaluation, confirm deep anesthesia by verifying the complete absence of pedal withdrawal reflex. Perform cervical dislocation as the secondary method of euthanasia, in accordance with AVMA Guidelines for the Euthanasia of Animals (2020 edition) and institutionally approved protocols.
   CAUTION: Animal carcasses, excised tissues, and blood-contaminated materials are biological/animal waste. Dispose of them according to institutional biosafety and animal facility regulations.
2. Perform a thoracotomy using surgical microscissors to expose the heart.
3. Insert a perfusion needle into the apex of the left ventricle and flush the systemic circulation with cold sterile phosphate-buffered saline (PBS). Successful perfusion is indicated by blanching of the liver and lungs.
4. Excise the intact heart and immerse it in 4% paraformaldehyde (PFA) at room temperature for a minimum of 24 h to ensure complete fixation.
   CAUTION: Paraformaldehyde is toxic and irritating. Handle 4% PFA in a chemical fume hood with appropriate PPE and dispose of PFA waste as hazardous chemical waste.
5. Transfer the fixed tissue to an automated tissue processor for sequential dehydration through a graded ethanol series, followed by xylene clearing and paraffin wax infiltration at 65 °C.
   CAUTION: Xylene is flammable and toxic, and hot paraffin can cause burns. Handle xylene in a fume hood or approved processor and dispose of xylene-containing waste as hazardous chemical waste.
6. Embed the processed hearts in paraffin blocks.
   NOTE: Correct heart orientation at embedding is critical. Position the heart to allow transverse sectioning through the aortic root at the level of the three valve leaflets.
7. Using a rotary microtome, cut serial cross-sections at 4–5 µm thickness through the aortic valve plane, ensuring that all three leaflets are captured. Mount sections on glass slides and store at room temperature.
8. Perform Hematoxylin and Eosin (H&E) staining on selected slides to assess tissue morphology, structural remodeling, and valve leaflet thickness.
9. Perform Von Kossa staining on adjacent serial sections to detect and quantify dystrophic calcification, identifiable as dark brown-to-black mineral deposits within the valve leaflets.

8. Sham surgical procedure

NOTE: This section describes the Sham control procedure. Sham mice undergo the same anesthesia, skin preparation, vessel exposure, arteriotomy, and retrograde guidewire insertion as the aortic wire injury (AWI) group; the guidewire is advanced to the supravalvular level but is not passed across the aortic valve, and no mechanical valve injury is performed.

1. Anesthetize the mouse and prepare the surgical field as described in steps 1.1. through 1.5.
2. Under stereomicroscopic visualization, expose the RCCA by blunt dissection as described in step 2.1.
3. Permanently ligate the distal end of the RCCA with a 6-0 silk suture and place a loose proximal slipknot (as in steps 2.2–2.3).
4. Perform the transverse arteriotomy and insert the guidewire as described in steps 3.1–4.2.
5. Advance the guidewire in a retrograde direction to the supravalvular level of the ascending aorta under echocardiographic guidance, confirming that the guidewire tip does NOT cross the aortic valve.
   NOTE: Do NOT perform the push-pull or rotational valve-injury motions described in step 4.4. Withdraw the guidewire and immediately ligate the proximal RCCA as described in steps 5.1–5.2.
6. Close the skin incision with interrupted 5-0 nylon sutures or wound clips.
   NOTE: As in the AWI group, intraoperative analgesia is provided by local lidocaine infiltration (step 1.5). Transfer the animal to a warmed recovery cage and monitor until fully ambulatory.
7. Subject Sham mice to identical post-operative monitoring, echocardiographic evaluation at 4 weeks, and histological processing as the AWI group.

9. Statistical analysis and study design

1. Across the study, 20 mice (10 male, 10 female) were subjected to the AWI procedure, and 6 C57BL/6 mice (3 male, 3 female) were subjected to the Sham procedure.
2. Measure peak transvalvular velocity (the criterion defining model success) in all surviving animals (19 AWI, 6 Sham).
3. Analyze detailed echocardiographic and histological indices in a pre-specified, sex-balanced subgroup of 6 AWI mice (3 male, 3 female).
4. Ensure all echocardiographic and histological measurements are performed by investigators blinded to group allocation.
5. Assess data for normality with the Shapiro-Wilk test before parametric testing. Compare two independent groups (AWI vs. Sham) using an unpaired two-tailed Student's t-test.
6. Analyze and display survival data using the Kaplan-Meier method.
7. Perform all analyses in GraphPad Prism. Present data as mean ± SEM, with p < 0.05 considered statistically significant.

Results

Development of a 3D-printed platform for hemorrhage control

To address perioperative hemorrhage—the principal source of morbidity and mortality in the conventional guidewire-induced AVS model—a 3D-printed vascular support platform was designed (Figure 1). The platform was fabricated by stereolithography (SLA) printing from a biocompatible (USP Class VI / ISO 10993-compliant) photopolymer resin, with a structural thickness of 1.07 mm. After printin...

Discussion

This protocol provides a detailed description of a stable, safe animal model of aortic stenosis. A feature of this approach is an auxiliary platform that facilitates guidewire insertion into the common carotid artery. The platform is fabricated by stereolithography (SLA) printing from a biocompatible photopolymer resin (structural thickness 1.07 mm) and incorporates an inverted V-shaped apical surface with a 120° apex angle; the complete design file is freely available as a supplementary .stp file. This device ensur...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
3D-printed vascular support platform	Custom-made	N/A	Custom 3D-printed vascular support platform; supplementary CAD file provided in .stp format.
4% Paraformaldehyde	Servicebio	G1101-3ML	Tissue fixation
5-0 nylon suture	Jinhuan Medical	F503	Skin closure by interrupted sutures
6-0 silk suture	Jinhuan Medical	RC411	Permanent ligation of distal RCCA and proximal slipknot
Acoustic coupling gel (pre-warmed)	Keppler	KL-250	Echocardiography
Biocompatible photopolymer resin	Formlabs	RS-F2-BMCL-01	SLA printing material used to fabricate the custom 3D-printed vascular support platform.
C57BL/6 mice (8–10 weeks; males 22–25 g, females 18–20 g)	GemPharmatech	C57BL/6	Both sexes used throughout; housed under standard conditions
Ethylene oxide gas	Institutional sterilization facility	N/A	Sterilization of the custom 3D-printed vascular support platform.
FreeCAD	FreeCAD Project Association	1.1.1	CAD visualization and generation of the 3D structural drawing from the .stp file.
GraphPad Prism	GraphPad Software	9.0	Statistical analysis and graphing
Guidewire, 0.014-inch diameter	Boston Scientific	H749393071900	Mechanical aortic valve injury
Hematoxylin and Eosin (H&E) staining kit	Beyotime	C0105S	Leaflet morphology and thickness
Isoflurane	RWD Life Science	R510-22-2	3% induction; 1.5–2% maintenance in 100% O2
Lidocaine	Shijiazhuang Pharmaceutical Group	H14024045	Pre-emptive local infiltration at incision site (0.5%, step 1.5)
Microsurgical scissors	RWD Life Science	F11022-11	Skin incision, arteriotomy, thoracotomy
MX550D transducer (40 MHz)	FUJIFILM VisualSonics	MX550D	Echocardiography
Nair hair removal cream	Nair	255 g	For anterior cervical and chest wall depilation
Phosphate-buffered saline (PBS)	Servicebio	G4202-100ML	Perfusion
Rotary microtome	Leica Biosystems	RM2125 RTS	Serial sectioning at 4–5 µm
Stereomicroscope (surgical)	Olympus	SZX7	Vessel isolation and microsurgery
Vevo 3100 high-resolution ultrasound system	FUJIFILM VisualSonics	Vevo 3100	Echocardiography
Von Kossa staining kit	Servicebio	G1043-20ML	Dystrophic calcification detection
Xylene	Nanjing Chemical Reagent Co., Ltd.	C0430530209	Tissue clearing during paraffin processing.