Establishment and Confirmation of a Postnatal Right Ventricular Volume Overload Mouse Model

Summary

This protocol presents the establishment and confirmation of a postnatal right ventricular volume overload (VO) model in mice with abdominal arteriovenous fistula (AVF), which can be applied to investigate how VO contributes to postnatal heart development.

Abstract

Right ventricular (RV) volume overload (VO) is common in children with congenital heart disease. In view of distinct developmental stages,the RV myocardium may respond differently to VO in children compared to adults. The present study aims to establish a postnatal RV VO model in mice using a modified abdominal arteriovenous fistula. To confirm the creation of VO and the following morphological and hemodynamic changes of the RV, abdominal ultrasound, echocardiography, and histochemical staining were performed for 3 months. As a result, the procedure in postnatal mice showed an acceptable survival and fistula success rate. In VO mice, the RV cavity was enlarged with a thickened free wall, and the stroke volume was increased by about 30%-40% within 2 months after surgery. Thereafter, the RV systolic pressure increased, corresponding pulmonary valve regurgitation was observed, and small pulmonary artery remodeling appeared. In conclusion, modified arteriovenous fistula (AVF) surgery is feasible to establish the RV VO model in postnatal mice. Considering the probability of fistula closure and elevated pulmonary artery resistance, abdominal ultrasound and echocardiography must be performed to confirm the model status before application.

Introduction

Right ventricular (RV) volume overload (VO) is common in children with congenital heart disease (CHD), which leads to pathological myocardial remodeling and a poor long-term prognosis^1,2,3. An in-depth understanding of RV remodeling and related early targeted interventions is essential for a good outcome in children with CHD. There are several differences in the molecular structures, physiological functions, and responses to stimuli in the hearts of adults and children^1,4,5,6. For example, under the influence of pressure overload, cardiomyocyte proliferation is the main response in neonatal hearts, whereas fibrosis occurs in adult hearts^5,6. In addition, many effective drugs in treating heart failure in adults have no therapeutic effect on heart failure in children, and may even cause further damage^7,8. Therefore, conclusions drawn from adult animals cannot be directly applied to young animals.

The arteriovenous fistula (AVF) model has been used to induce chronic heart VO and corresponding cardiac dysfunction for decades in adult animals of different species^{9,10,11,12,13}. However, little is known about the model in postnatal mice. In our previous studies, a VO postnatal mouse model was successfully generated by the creation of an abdominal AVF. The changed RV developmental track in the postnatal heart was also demonstrated^14,15,16,17.

To explore the underlying modified surgical process and characteristics of the present model, a detailed protocol is presented; the model is evaluated for 3 months in this study.

Protocol

All of the procedures presented here conformed to the principles outlined in the Declaration of Helsinki and were approved by the Animal Welfare and Human Studies Committee at Shanghai Children's Medical Center (SCMC-LAWEC-2023-003). C57BL/6 mice pups (P7, males, 3-4 g) were used for the present study. The animals were obtained from a commercial source (see Table of Materials). The mice pups and their nursing mothers (pups:mothers = 6:1 in a single cage) were kept under specific-pathogen-free laboratory conditions under a 12 h light and dark cycle at 22 ± 2 °C with free access to water and a nutritional diet. The pups were randomized into two groups: a VO group and a sham-operated (sham) group.

1. Equipment and surgical tool preparation

NOTE: The commercial details of all the materials/equipment are listed in the Table of Materials.

1. Ensure that the following types of equipment are ready and properly functioning: operating table (foam plastic panel), inhalational anesthesia machine, microscope with vertical illumination and a built-in camera, ultrasound device with a 24 MHz transducer, and thermostatic heating platform.
2. Sterilize the surgical instruments (i.e., a micro-needle holder, fine-tip forceps, and round-handled Vannas spring scissors).
3. Assemble the following consumables: 11-0 and 9-0 surgical suture needles (taper-point) with thread, tape strips, 5 mL syringe needles, 2-0 silk (surgical fixation), sterile cotton swabs, and ultrasound gel.
4. Ensure the following reagents are present: Betadine, 70% ethanol, normal sterile saline, isoflurane, acetaminophen, ophthalmic ointment, and hair removal cream.

2. Surgical procedure

NOTE: The fistula surgery procedure was modified according to the previously described method¹¹. Figure 1 shows a schematic diagram of the AVF operation in postnatal mice.

1. Anesthesia and restraint
   1. Place the mice pups into an anesthesia-induction box supplied with 2% isoflurane/oxygen for 2 min with the flow set to 1 L/min. Administer acetaminophen (0.1 ml PO of 80 mg/2.5 ml) using a TB syringe.
   2. Place the pups in a supine position on the operating table with nasal inhalation of 1.5% isoflurane with a 0.8 L/min flow to maintain anesthesia. Adjust the pup's position by tying the legs to the fixed syringe needles. Apply ophthalmic ointment to the pups' eyes to prevent corneal desiccation.
   3. Pinch the anesthetized pup's tail to check its pain responsiveness; no obvious body movements indicate adequate anesthesia.
2. Fistula surgery
   1. Disinfect the skin with three alternating scrubs of betadine and 70% ethanol, and then drape the surgical site. Cut the abdominal wall and peritoneum from the lower abdomen to the subxiphoid to fully expose the peritoneal cavity, taking care not to injure the abdominal organs. Drip normal sterile saline to moisten externalized organs.
   2. Gently pull the gastrointestinal tract and bladder away from the surgical site using cotton swabs to visualize the vertical abdominal aorta (AA) and inferior vena cava (IVC) under the retroperitoneum. Rotate the operating table 90° counterclockwise and adjust the microscope magnification to visualize the two horizontal vessels clearly.
   3. Puncture the fistula from the AA into the IVC in an oblique direction 1 cm distal to the renal artery with an 11-0 suture needle (diameter = 0.07 mm). Verify successful fistula creation based on the swelling and mixing of venous and arterial blood in the IVC.
   4. Next, rapidly compress the bleeding point using an appropriate force applied with dry cotton swabs for 15 s. Replace the stomach, intestines, and bladder in the abdominal cavity as soon as possible to promote hemostatic compression.
   5. Suture the abdominal wall and peritoneum with a blanket stitch using a 9-0 suture thread. Discontinue anesthesia and provide the pups with 100% oxygen for 1 min.
3. Anesthesia resuscitation
   1. Place the pups on a 38 °C heating platform. After a complete awakening with vitality, return the pups to their nursing mother. The entire procedure lasts approximately 15 min.
      ​NOTE: In the present study, the sham group undergoes the same procedure except for the puncturing step.

3. Ultrasound confirmation of fistula

NOTE: The general operation of the ultrasound device was identical to previous reports^18,19.

1. Confirmation of fistula by abdominal ultrasound
   1. After induction of anesthesia (step 2.1.1), fix the mice with tape strips in the supine position on the warm platform. Then, connect the mice to an electrocardiogram (ECG) monitor with ultrasound gel. Maintain anesthesia using 1.5% isoflurane at a 0.8 L/min flow.
   2. Prepare the chest and abdominal skin using hair removal cream. After a few seconds, remove the cream with a warm water-soaked cotton tip. Place the transducer (24 MHz) on the midabdominal line and rotate the transducer marker to the head of the mice.
   3. Move the platform down to the left or right side of the mice and use B-mode and color Doppler mode to visualize the long-axis view of the vessels and blood signals^18,19. Measure the blood flow velocity of the AA, IVC, and fistula to confirm AVF patency through pulsed wave Doppler mode.
      NOTE: Successful fistula creation on the ultrasound was indicated by a turbulent flow signal visible between the AA and IVC (Figure 2C). Doppler blood flow velocity at the AVF site was significantly elevated compared with a relatively lower systolic velocity in the AA (Figure 2A,C). Moreover, in contrast with normal flow patterns in the IVC (Figure 2B), the pulsatile waveform of IVC blood flow proximal to the AVF also confirmed the successful creation of the fistula (Figure 2D).
2. Confirmation of VO by echocardiography
   1. Move the tail-end part of the platform downward, place the transducer (24 MHz) on the chest, and rotate the transducer marker to the right shoulder of the mice. Visualize the modified parasternal long-axis view of the pulmonary artery (PA) using B-mode and color Doppler mode.
   2. Using pulsed wave Doppler mode, measure blood flow signals in the PA, including the velocity time integral (PA-VTI), diameter of the PA valve (PAD), pulmonary arterial acceleration time (PAT), and RV ejection time (RVET) (Figure 2E,F and Figure 3A,B).
   3. Measure the ultrasound parameters from the mean of three consecutive measurements. Calculate the RV stroke volume (RVSV, mL) and RV systolic pressure (RVSP, mmHg) using the following formulas²⁰:
      RVSV [mL] =1/4 × πD² × VTI_PA
      RVSP [mmHg] = -83.7 × PAT/RVET - index + 63.7
      NOTE: Considering the ultrasound measurement bias, an increase of >15% in the RVSV or VTI_PA in VO mice compared to mice in the sham group was considered VO in the RV (Figure 2E,F).

Results

Survival rate and AVF patency within 3 months
A total of 30 (75%) mice in the VO group and 19 (95%) mice in the sham group survived the AVF surgery (Figure 4A). In the VO group, eight mice died within 1 day after surgery due to excessive bleeding (n = 5) or cannibalization (n = 3), whereas two mice died of unknown causes at 1 month.

Of the surviving VO mice (n = 30), ultrasound confirmed the successful establishment of fistulas in 21 mice post...

Discussion

Previously, the classic RV VO model was created using valve regurgitation²¹; however, compared to AVF, open-heart valve surgery may require more sophisticated techniques and may be associated with significantly higher mortality, particularly in postnatal mice. As animal studies have shown that the same effect of VO has been achieved by AVF²², modified abdominal fistula surgery with less trauma was used in this study.

Certain factors were consider...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
70% Ethanol	Tiandz,Chia
ACETAMINOPHEN Oral Solution	VistaPharm, Inc. Largo, FL 33771, USA	NDC 66689-054-01
Anesthesia machine	RWD Life Science,China	R550IP
Anesthesia mask	RWD Life Science,China	68680
C57BL/6 mice	Xipu’er-bikai Experimental Animal Co., Ltd (Shanghai, China)
Hair removal cream	Veet, France	VT-200
Hematoxylin and eosin Kit	Beyotime biotech	C0105M
Isoflurane	RWD Life Science,China	R510-22-10
Microscope	Yuyan Instruments, China	SM-301
Surgical suture needles	NINGBO MEDICAL NEEDLE CO.,LTD, China
Thermostatic heating platform	Qingdao Juchuang Environmental Protection Group Co., Ltd, China
Ultrasound device	FUJIFILM VisualSonics, Inc.	Vevo 2100	Image modes includes B-Mode, Color Doppler Mode and Pulsed Wave Doppler Mode
Ultrasound gel	Parker Laboratories,United States	REF 01-08
Ultrasound transducer	FUJIFILM VisualSonics, Inc.	MS 400