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

Published: June 9, 2023 doi: 10.3791/65372
* These authors contributed equally


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.


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.


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 prognosis1,2,3. 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 children1,4,5,6. For example, under the influence of pressure overload, cardiomyocyte proliferation is the main response in neonatal hearts, whereas fibrosis occurs in adult hearts5,6. 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 damage7,8. 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 species9,10,11,12,13. 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 demonstrated14,15,16,17.

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.

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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 reports18,19.

  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 signals18,19. 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 × VTIPA
      RVSP [mmHg] = -83.7 × PAT/RVET - index + 63.7
      NOTE: Considering the ultrasound measurement bias, an increase of >15% in the RVSV or VTIPA in VO mice compared to mice in the sham group was considered VO in the RV (Figure 2E,F).

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Representative 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 postoperatively, which were shown to be patent at 1 week postoperatively (P14) and maintained until 2 weeks postoperatively (P21). However, the fistula closed at 1 month in seven mice and at 2 months in two mice. Only 12 mice had a persistent AVF at the 3 month follow-up. The AVF patency rates were 70, 70, 46.7, and 40% at 1 week, 2 weeks, 1 month, and 2 months postoperatively, respectively (Figure 4B).

Hemodynamic changes in the right heart
The 3-month follow-up of the hemodynamic parameters showed that both the PAD and RVSV of the mice in each group increased with age within two months (n = 6 in both groups; Figure 3B,E,F). Compared to the sham-operated mice, PA-VTI was significantly higher in the VO group within 2 weeks postoperatively (Figure 3D) but declined thereafter, and PA flow patterns changed with decreasing PAT (Figure 3A). The RVSV in the VO group was consistently higher than that in the sham group for 2 months, with an increase of approximately 30%-40%. The RVSP was significantly increased with pulmonary regurgitation 2 months after surgery (Figure 3C,G).

Morphological changes in the right heart and small pulmonary arteries
Under the microscope, the RV was significantly enlarged compared to the sham group after the AVF (Figure 5A). Histological staining showed a thickened RV-free wall and enlarged RV cavity in the VO mice (Figure 5B). According to the RV hemodynamic changes, the RVSP was elevated 2 months after surgery. Lung tissues from two groups of mice 3 months after surgery were randomly selected for hematoxylin and eosin (HE) staining, which showed a thickened tunica media, endothelial hyperplasia, and peripheral inflammatory cell infiltration in some of the small pulmonary arteries of the VO group (Figure 5C).

Figure 1
Figure 1: Schematic diagram of the AVF operation in postnatal mice. (A) Surgical instruments. (B) The procedure of AVF surgery. Abbreviations: AVF = arteriovenous fistula; IVC = inferior vena cava; AA = abdominal aorta. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Confirmation of AVF fistula and VO by ultrasound. (A) Normal pulsatile flow signal in the AA (peak flow velocity: 400 mm/s). (B) Normal blood flow signal of the IVC. (C) Increased flow velocity at the fistula (the red blood flow signal with a yellow and green hue inside indicated a turbulent flow signal at the fistula; peak systolic flow velocity: 900 mm/s). (D) Pulsatile flow in the IVC near the fistula with the increased flow velocity. (E) Increased PA-VTI in VO mice 1 week after surgery. (F) PA-VTI in sham mice 1 week after surgery (the blue blood flow signal indicated the blood flow of the PA). Abbreviations: AVF = arteriovenous fistula; IVC = inferior vena cava; AA = abdominal aorta; PA = pulmonary artery; VTI = velocity-time integral. In Doppler color mode, flow toward the transducer was encoded in red, and away from the transducer was encoded in blue. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Hemodynamic measurements of the right heart derived from echocardiography. (A) Doppler flow patterns at each time point of VO mice showed gradually decreasing PAT. (B) Two-dimensional measurements of PA parameters. (C) PA regurgitation on color Doppler echocardiography. (D-F) Changes in PA-VTI, PAD and RVSV at each time point in postoperative VO mice. (G) Histogram of RVSP in VO (black) and sham (Gray) mice showed an increased RVSP in 2 months and 3 months after AVF surgery (six VO mice; six sham mice; Student's t-test; *represents statistical significance). Abbreviations: P14 = postnatal day 14; P21 = postnatal day 21; PVR = pulmonary valve regurgitation; RVSP = right ventricular systolic pressure; M = months; W = weeks. The figure F is adapted from Sun et al. with permission14. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Survival rate and fistula patency rate of mice after AVF surgery. (A) The survival rate of postnatal mice after surgery (n = 40 in the VO group; n = 20 in the sham group). (B) Fistula patency rate in VO mice (n = 30). Please click here to view a larger version of this figure.

Figure 5
Figure 5: Morphological changes in the right heart. (A) Enlarged heart in VO mice at each time point after AVF surgery. (B) Cardiac HE staining at different time points after surgery showed a thickened RV-free wall and enlarged RV cavity. (C) Histopathological changes of pulmonary arterioles in mice after AVF showed hyperplasia and hypertrophy of small pulmonary arteries with infiltration of inflammatory cells. Scale bars: (A) = 5 mm; (B) = 2000 µm; (C) = 50 µm. Abbreviations: W = weeks; M = months. Please click here to view a larger version of this figure.

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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 considered during the procedure to successfully establish the fistula. First, the procedure was conducted in postnatal mice without endotracheal intubation and assistant ventilation; therefore, prompt adjustment of the anesthesia settings for the pups according to their dynamic condition was essential to avoid death from respiratory failure. Second, the pup's stomach and bladder were frequently in a full state during surgery. Therefore, to adequately expose the retroperitoneal vascular structures, a gentle, delicate operation was needed to avoid injury to the fragile abdominal organs. Third, ligation of the AA to prevent severe arterial bleeding was difficult to perform in the pups; therefore, hemostatic compression using cotton swabs immediately after the puncture was required. Thereafter, it was possible for the retroperitoneum and abdominal organs to produce further compression on the bleeding site. Moreover, it was noted that excessive compression may contribute to early fistula failure.

Owing to the less traumatic procedure compared to adult RV VO models, postnatal VO mice showed a relatively higher perioperative survival but lower fistula success rates in the early postoperative period11,23. In addition to severe hemorrhage, cannibalizing the inexperienced mother was the main cause of death in the pups after surgery. A comfortable and quiet breeding environment, tight closure of abdominal wounds, rapid body temperature recovery, and full awakening of the pups after anesthesia may reduce the risk of cannibalization. Previous studies on adult AVF mouse models have found that the formation of AVF has three stages: rapid thrombosis period on postoperative days 0-1, fistula maturity period for 3 weeks, and finally successful AVF creation with fistula reclosure in a few mice in 3-6 weeks23. In this study, the fistula patency curve of the postnatal mice also showed the same trajectory (i.e., fistula closure mainly occurred within 1 week or during 4-8 weeks after surgery, and the remaining fistulae remained open at 3 months). Therefore, it is critical to confirm the patency of the fistula in the postnatal AVF mice within 2 months after surgery by abdominal ultrasound.

An increased RVSV is another essential piece of evidence for RV VO, except for fistula patency. At present, cardiac catheterization is difficult to implement in low-weight, young mice. Benefitting from the advantages of its noninvasiveness, relatively simple manipulation, and continuous monitoring of the same mouse, echocardiography with high-frequency transducers was applied to evaluate the hemodynamic changes in this study. RVSV was estimated by pulmonary blood flow VTI, and it increased by about 30%-40% within 2 months after surgery in the postnatal VO mice. These results further proved the successful establishment of AVF and RV VO in this model.

Chronic VO may gradually lead to functionally elevated pulmonary resistance and finally vascular remodeling of PA arterioles. This process is common in children with CHD with a left-to-right shunt. Previous animal studies in sheep and piglets have proved that AVF could lead to structural and functional changes in the pulmonary vascular system13,24,25. During the subsequent follow-up at 2 months after surgery, abnormal morphological patterns of PA Doppler flow with decreased PAT, pulmonary valve regurgitation, and a downward trend of RVSV in VO mice were observed. As previously reported, PAT can be used as a complementary parameter to evaluate the RV afterload in neonates and children. The phenomena mentioned above may suggest an altered pulmonary vascular resistance in the VO mice26,27,28. To quantify the elevated RV afterload or pressure overload, the ratio of PAT and RVET was used to estimate the value of RVSP using the formula verified by Thibault in adult mouse18, which demonstrated that RVSP was significantly increased 2 months after AVF surgery in the postnatal model. In addition, histopathological evidence of inflammation and PA remodeling in several lung lobes of VO mice further proved the structural abnormalities 3 months after surgery. Therefore, to exclude the effect of pressure overload, it was suggested that the application of this postnatal mice RV VO model was limited to 2 months after surgery.

In summary, modified AVF surgery is a feasible technique to establish the RV VO model in postnatal mice. Considering the probability of fistula closure and elevated pulmonary artery resistance, abdominal ultrasound, and echocardiography should be performed to confirm the model status before application.

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There are no conflicts of interest to declare.


This work was supported by the National Science Foundation of China (no. 82200309) and the Innovation Project of Distinguished Medical Team in Ningbo (no. 2022020405)


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



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Postnatal Right Ventricular Volume Overload Mouse Model Congenital Heart Disease Developmental Stages Myocardium Abdominal Arteriovenous Fistula Morphological Changes Hemodynamic Changes Abdominal Ultrasound Echocardiography Histochemical Staining Survival Rate Fistula Success Rate RV Cavity Enlargement Thickened Free Wall Stroke Volume Increase RV Systolic Pressure Increase Pulmonary Valve Regurgitation Pulmonary Artery Remodeling Arteriovenous Fistula Surgery Model Status Confirmation
Establishment and Confirmation of a Postnatal Right Ventricular Volume Overload Mouse Model
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Sun, S., Zhu, H., Wang, S., Xu, X.,More

Sun, S., Zhu, H., Wang, S., Xu, X., Ye, L. Establishment and Confirmation of a Postnatal Right Ventricular Volume Overload Mouse Model. J. Vis. Exp. (196), e65372, doi:10.3791/65372 (2023).

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