Surgically Induced Cardiac Volume Overload by Aortic Regurgitation in Mouse

Summary

This protocol presents a practical guide on the surgery for creation of aortic regurgitation (AR) in the mouse. Assessment of the AR mouse by echocardiography and invasive hemodynamic measurement recapitulates its clinically relevant characteristics of volume overload-induced eccentric hypertrophy, suggesting its promising application in the study of cardiac hypertrophy.

Abstract

Aortic regurgitation (AR) is a common valvular heart disease that exerts volume overload on the heart and represents a global public health problem. Although mice are widely applied to shed light on the mechanisms of cardiovascular disease, mouse models of AR, especially those induced by surgery, are still paucity. Here, a mouse model of AR was described in detail which is surgically induced by disruption of the aortic valves under high-resolution echocardiography. In accordance with regurgitated blood flow, AR mouse hearts present a distinctive and clinically relevant volume overload phenotype, which is characterized by eccentric hypertrophy and cardiac dysfunction, as evidenced by echocardiographic and invasive hemodynamic evaluation. Our proposal, in a reliable and reproducible manner, provides a practical guide on the establishment and assessment of a mouse model of AR for future studies on molecular mechanisms and therapeutic targets of volume overload cardiomyopathy.

Introduction

In the presence of increased volume overload (preload) or pressure overload (afterload), the heart undergoes enlargement, a condition termed hypertrophy. Although cardiac hypertrophy is a compensatory response to maintain perfusion of peripheral organs before cardiac failure, it is also an independent risk factor for major cardiovascular events^1,2. Volume overload is one of the important manifestations of increased mechanical stress. Volume overload occurs during cardiac diastole and induces eccentric cardiac hypertrophy, which is not only commonly seen in valvular diseases, such as aortic regurgitation and mitral regurgitation, but also in end-stage hypertensive heart disease, myocardial infarction, dilated cardiomyopathy, and excessive exercise. In addition, in clinical practice, some drugs that can better reduce the myocardial hypertrophy induced by pressure overload have unsatisfactory effects in the treatment of myocardial hypertrophy induced by volume overload¹. It is therefore of great significance to discover the mechanism and intervention methods of eccentric cardiac remodeling caused by volume overload. However, such research on volume overload has been significantly hampered for a long time, which can be, in large part, attributed to the lack of small animal models that can be easily operated, efficiently quantified, and stably replicated³.

As for small animal species, mice have become the mainstream model animal for cardiovascular disease research due to their short life cycle, convenient operation, clear genome, and ease of genetic modification⁴. In terms of model categories, compared to genetic modification models and drug-treated models, surgical models have obvious unique advantages. The surgical model can avoid excessive and laborious mouse breeding and gene identification that are necessary for the genetic modification model and can also avoid the non-specific effects on extracardiac tissues and organs that are difficult to control in drug-treated models. The mouse model of aortocaval shunt has been documented to induce cardiac volume overload in previous literature⁵. However, aortocaval shunt accounts for a small fraction of cardiac eccentric hypertrophy in the clinic and causes biventricular overload⁵, making it of little translational significance to be used in left ventricular eccentrical hypertrophy study. Nevertheless, valvular heart disease represents a major public health problem worldwide; it is estimated that around 15% of the population >75 years of age has significant valvular disorder⁶. Although aortic regurgitation (AR) occupies a portion of valvular heart disease, it distinctively causes eccentric left ventricular (LV) hypertrophy due to an increase in volume overload by regurgitant blood flow^7,8. Considering the right common carotid artery (RCCA) provides a route to reach the location of aortic valves, it is conceptually intriguing to disrupt aortic valves via the RCCA to cause regurgitant blood flow in mice. Inspired by the techniques of creating oscillating aortic flow⁹, a mouse model of aortic regurgitation (AR) was recently established in our lab to surgically induce volume overload⁷. This AR mouse demonstrates obvious LV eccentric hypertrophy, which is a clinically transformative approach and demonstrates a great translational potential for studying the overloaded heart phenotype and its underlying mechanism. Here, a detailed step-by-step procedure was described to perform AR surgery in mice, recapitulated by high frequency echocardiography and invasive hemodynamics to ensure the success of the surgery (Figure 1).

Protocol

This protocol has received ethical approval from the Animal Care and Use Committee of Zhongshan Hospital, Fudan University, and follows the recommendations of Guide for the Care and Use of Laboratory Animals (No. 85-23, revised 2011; National Institutes of Health, Bethesda, MD, USA).

NOTE: Animal experiments were performed on male C57BL/6J mice >10 weeks of age. The surgeon in this protocol should be skillful in the manipulation of murine echocardiography, before he/she performs the AR operation in the mouse. However, at most research institutions, small rodent echocardiography is operated by a core facility, so the surgeon can closely collaborate with core experts, if not an experienced surgeon in echocardiography. Experience of invasive hemodynamic measure in mouse is a plus.

1. Preparation for ultrasound imaging (mandatory) and invasive hemodynamic measurement (optional)

1. Start up the ultrasound machine connected to a 30 MHz probe. Set the temperature-controlled ultrasound animal platform in the position for the aortic arch view, in which the right side of the mouse is tilted up.
   NOTE: It is recommended that the cranial end of the ultrasound animal platform is placed toward the surgeon. However, whether the cranial end or the caudal end is toward the surgeon should be dependent upon on which one the surgeon feels more comfortable with.
2. Connect a micromanometer (pressure catheter) to the data acquisition device and analog/digital converter. Immerse the micromanometer's calibration cuvette in saline for saline calibration.
   ​NOTE: If the condition permits, a pressure-volume catheter can be used as well. We use a pressure catheter because the pressure data acquisition device in the lab collects pressure-only data and does not have the capacity to collect volume-related data, although the echocardiographic results in the current study can also delineate LV volumes.

2. Anesthesia of mice, preparation of surgical devices, and isolation of the RCCA

NOTE: Surgical tools must be sterilized and autoclaved before use. All steps are recommended to be performed under aseptic conditions. It is also recommended that hair removal is performed 1 day ahead to save time during the imaging procedure, minimize potential undesired stress responses in the mice, and to keep the chest and extremities clean and dry.

1. Anesthetize the mouse in the induction chamber, which is connected to a vaporizer set to 4% isoflurane mixed with 0.8 L/min of oxygen. When the mouse falls asleep or the tail pinch reflex disappears, remove the animal from the induction chamber.
2. Place the animal in the supine position on a copper plate, which is warmed by a heating pad. Connect its nose to a nosecone, to which 1.5% isoflurane mixed with 0.8 L/min of oxygen is delivered for maintaining a steady level of anesthesia.
   NOTE: A copper plate is recommended as it is convenient to clean and is rust proof, though it can be replaced by another type of metal plate.
3. Place ophthalmic ointment on the eyes to prevent dryness under anesthesia and tape the extremities onto the copper plate. Remove hair from the neck and chest using depilatory cream and clean the depilated area with 75% ethanol.
4. Prepare the necessary surgical tools, including various forceps and scissors (Figure 2A; see Table of Materials).
5. Make a longitudinal median incision, around 1 cm, in the neck with curved thumb forceps and straight scissors, between the lower jaw and sternum.
6. Bluntly dissect the left and right part of the thyroid gland using two pairs of forceps. With the curved fine thumb forceps, separate the stemohyoideus muscle and fat tissue in the right paratracheal region to expose the RCCA for as long as possible. Avoid injury of the vagal nerve at all times, as this can cause hypotension, bradycardia, and death (Figure 2B).

3. Catheterization through the RCCA and ascending aorta under ultrasound guidance

1. Pass two 6-0 silk threads, around 5 cm each, under the vessel. Ligate the distal RCCA with a tight knot using one thread and fix the two ends of the tight knot next to the head of the animal to maintain light tension on the RCCA. This action will facilitate the catheterization in the coming steps.
2. Place a loose knot on the proximal RCCA using the second thread. This fills the sealed region of the RCCA with blood, making it easy to incise.
3. Use small pinch scissors to cut a wedge-shaped opening, 1-2 mm proximal to the tight knot, to open the RCCA. Make sure the size of the incision is neither too small to insert a catheter, nor too large for it to snap during insertion.
   NOTE: Incision under a microscope is highly recommended. Puncturing a small hole in the vessel with a 26 G needle is an alternative method.
4. Prepare a plastic catheter containing a metal wire (Figure 2C). Stretch the incision with long-handed curved tying forceps, insert the plastic catheter containing the metal wire into the RCCA, and move forward to the loose knot.
5. Relieve the loose knot to advance the catheter and wire around 2 cm. Transfer the copper plate containing the animal onto the ultrasound animal platform, apply ultrasound gel to the mouse neck and chest, and then carefully forward the catheter and wire through the RCCA and ascending aorta under ultrasound guidance.

4. Puncture of the aortic valves under ultrasound guidance

1. Collect basal ultrasound data in color Doppler mode and pulse wave Doppler mode before the plastic catheter and metal wire reach the aortic orifice.
2. With the ultrasound simultaneously and clearly showing the ascending aorta, the LV outflow tract, the catheter, and the wire, when the catheter and wire reach the aortic orifice, protrude the tip of the wire from the catheter, and puncture the aortic valves (Figure 1).
   NOTE: When the aortic valve is perforated, the surgeon should be able to sense this break.
3. Slightly retreat the catheter and wire from the aortic orifice and collect post-perforation ultrasound data in color Doppler mode and pulse wave Doppler mode after puncture of the aortic valves. The regurgitant flow is red in color during cardiac diastole in color Doppler mode and can be quantitively confirmed in pulse wave Doppler mode.
4. Consider a peak diastolic velocity of aortic flow (PSVa) between 300-500 mm/s as satisfactory. If the regurgitant degree of blood flow is unsatisfactory, repeat step 4.2.
5. Optional: Apply a micromanometer before and instantly after perforation of the aortic valves to further confirm the existence of regurgitant flow. To check, both the aortic end-diastolic pressure (AEDP) is depressed, and the aortic pulse pressure is enhanced by around 20 mmHg.
   ​NOTE: A detailed description of how to use a micromanometer catheter to perform invasive LV hemodynamic measurement has been elegantly presented elsewhere^10,11.

5. Withdrawal of the plastic catheter and metal wire, and perioperative care

1. Remove the ultrasound gel and dry the mouse with sterile gauze or tissue after confirmation of successful perforation of the aortic valves, then carefully withdraw the plastic catheter with the central metal wire, before ligation of the RCCA.
2. Close the skin using a 5-0 silk suture in a continuous suture pattern and apply povidone-iodine solution to the suture site. Administer the mouse with meloxicam (0.13 mg) subcutaneously for analgesia and place the mouse in a pre-warmed cage under a warming light until fully awake for recovery.

6. Sham surgery

1. Perform sections 1-3 as described. For the sham-operated mouse, perform similar procedures as in section 4 without disruption of the aortic valves.
2. Perform section 5 as described, although AR should not be present in any of the sham-operated mice.

7. Assessment of aortic valve perforation, cardiac morphology, and function using echocardiography and invasive hemodynamic measurement

1. After 4 weeks of AR, use echocardiographic B-mode, color Doppler mode, and pulse wave Doppler mode to assess blood flow of the aortic arch in the aortic arch view, and measure PDVa, according to step 4.1 and elsewhere^1,12.
2. Use echocardiographic B-mode and M-mode to assess LV dimension and contractility in the parasternal long axis view, with the LV end-diastolic (LVEDD) and end-systolic (LVESD) dimensions, LV posterior wall end-diastolic (LVPWTd) and end-systolic (LVPWTs) thickness, LV ejection fraction (LVEF), and fractional shortening (LVFS) derived.
   NOTE: A detailed description of how to use the ultrasound machine and manipulation of ultrasound views has been elegantly described previously¹².
3. After the echocardiographic imaging, perform invasive hemodynamic measurement, in a manner similar to step 4.3 and elsewhere¹⁰. Record the maximal contraction and relaxation velocity (+dp/dt and −dp/dt). Insert the micromanometer into the left common carotid artery (LCCA, not RCCA), since the RCCA was permanently ligated during the AR surgery.
4. After the invasive hemodynamic measurement, euthanize the mouse via cervical dislocation. Open the chest, flush the heart with 10% formalin, followed by 0.9% sodium chloride solution, excise the heart by cutting off the aorta, and section transversely at the level of the inferior margin of the left auricle. Acquire images using light microscopy.

Results

To guarantee successful AR, we validated regurgitant blood flow using color Doppler and pulse wave Doppler echocardiography. In mice with AR, the color Doppler spectrum of the aortic arch showed regurgitant flow (red) immediately post-operation, which was absent in sham mice (no flow in diastole; Figure 3A). Consistently, the pulse wave Doppler demonstrated robustly elevated regurgitant flow in AR mice (Figure 3B,C). With a further confirmation ...

Discussion

The surgical induction of AR in the mouse is a technically challenging, new technique but has significant translational relevance. To master the technique, a surgeon should at least be familiar in advance with murine cervical and cardiac anatomy, mouse handling, and echocardiography. Skillful operation in invasive hemodynamic measurement is a plus. For successful AR operation, special care should be taken on several critical steps.

Cutting open the RCCA is the most crucial step. The hole on th...

Materials

Name	Company	Catalog Number	Comments
Copper plate	JD.com Inc.	Customized	20 X 15 cm or bigger is prefeered
Curved Tying forceps	66 Vision Tech	53324A	to stretch and isolate muscle, tissue, and vessel
Heating pad	JD.com Inc.	Changzhi 55	warm the copper plate and mouse by the way
Long-handed Curved Tying Forceps	MECHENIC	TS-15	to stretch vessel
Metal Wire (stainless steel)	JD.com Inc.	0.18 mm in diametter	work with a plastic catheter to puncture aortic valves
Needle Holder	Shanghai Jinzhong	131110	suture of skin
Plastic Catheter	Anilab software & instruments	PE-0402	work with a metal wire to puncture aortic valves
Pressure Catheter	Millar Instruments	SPR 835	1.4F in size
Pressure Data Acquisition Device and Analog/Digital Converter	AD Instruments	Labchart 5	connected with pressure catherter
Scissor	Suzhou Shiqiang	Stronger 13Cr	to cut skin
Smallpinch Scissors	Shanghai Jinzhong	YBE030	to cut vessel
Stereomicroscope	Olympus Corporation	SMZ845	for incision and intubation of vessel
Straight Tying forceps	66 Vision Tech	53320A	to stretch and isolate muscle, tissue, and vessel
Thumbforceps	Suzhou Shiqiang	5307B	to clamp and stretch skin and muscle
Ultrasound Gel	PARKER	Aquasonic-100	to transfer ultrasound signal
Ultrasound Imaging System	VisualSonics	2100	includes B-mode, M-model, color Doppler and pulse wave Dopper
Vaporizer	RWD Life Science	R540	for anesthesia