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)

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&#39;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 &#177; 2 &#176;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.
<ol>
	<li>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.</li>
	<li>Sterilize the surgical instruments (i.e., a micro-needle holder, fine-tip forceps, and round-handled Vannas spring scissors).</li>
	<li>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.</li>
	<li>Ensure the following reagents are present: Betadine, 70% ethanol, normal sterile saline, isoflurane, acetaminophen, ophthalmic ointment, and hair removal cream.</li>
</ol>
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.
<ol>
	<li>Anesthesia and restraint
	<ol>
		<li>Place the mice pups into an anesthesia-induction box supplied with 2% isoflurane/oxygen for 2 min&#160;with the flow set to 1 L/min. Administer acetaminophen (0.1 ml PO of 80 mg/2.5 ml) using a TB syringe.</li>
		<li>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&#39;s position by tying the legs to the fixed syringe needles. Apply ophthalmic ointment to the pups&#39; eyes to prevent corneal desiccation.</li>
		<li>Pinch the anesthetized pup&#39;s tail to check its pain responsiveness; no obvious body movements indicate adequate anesthesia.</li>
	</ol>
	</li>
	<li>Fistula surgery
	<ol>
		<li>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.</li>
		<li>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&#176; counterclockwise and adjust the microscope magnification to visualize the two horizontal vessels clearly.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Anesthesia resuscitation
	<ol>
		<li>Place the pups on a 38 &#176;C heating platform. After a complete awakening with vitality, return the pups to their nursing mother. The entire procedure lasts approximately 15 min. 
		&#8203;NOTE: In the present study, the sham group undergoes the same procedure except for the puncturing step.</li>
	</ol>
	</li>
</ol>
3. Ultrasound confirmation of fistula
NOTE: The general operation of the ultrasound device was identical to previous reports18,19.
<ol>
	<li>Confirmation of fistula by abdominal ultrasound
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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).</li>
	</ol>
	</li>
	<li>Confirmation of VO by echocardiography
	<ol>
		<li>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.</li>
		<li>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).</li>
		<li>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&#160;&#215;&#160;&#960;D2 &#215; VTIPA 
		RVSP [mmHg]&#160;= -83.7 &#215; PAT/RVET - index + 63.7 
		NOTE: Considering the ultrasound measurement bias, an increase of &#62;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).</li>
	</ol>
	</li>
</ol>

Modified Arteriovenous Fistula Surgery to Establish the RV Volume Overload Mouse Model

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The right heart in adults with congenital heart disease.</a> Revista Espa&#241;ola de Cardiolog&#237;a. 63 (9), 1070-1086 (2010).</li><li>Kovacs, A., Lakatos, B., Tokodi, M., Merkely, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Right+ventricular+mechanical+pattern+in+health+and+disease:+beyond+longitudinal+shortening.">Right ventricular mechanical pattern in health and disease: beyond longitudinal shortening.</a> Heart Failure Reviews. 24 (4), 511-520 (2019).</li><li>Ye, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+blood+oxygen+saturation+during+postnatal+human+cardiomyocyte+cell+cycle+activities.">Role of blood oxygen saturation during postnatal human cardiomyocyte cell cycle activities.</a> JACC: Basic to Translational Science. 5 (5), 447-460 (2020).</li><li>Ye, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pressure+overload+greatly+promotes+neonatal+right+ventricular+cardiomyocyte+proliferation:+a+new+model+for+the+study+of+heart+regeneration.">Pressure overload greatly promotes neonatal right ventricular cardiomyocyte proliferation: a new model for the study of heart regeneration.</a> Journal of the American Heart Association. 9 (11), e015574 (2020).</li><li>Geraets, I. 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There are no conflicts of interest to declare.

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...

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.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>70% Ethanol</td>
			<td>Tiandz,Chia</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ACETAMINOPHEN Oral Solution</td>
			<td>VistaPharm, Inc. Largo, FL 33771, USA</td>
			<td>NDC 66689-054-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Anesthesia machine</td>
			<td>RWD Life Science,China</td>
			<td>R550IP</td>
			<td></td>
		</tr>
		<tr>
			<td>Anesthesia mask</td>
			<td>RWD Life Science,China</td>
			<td>68680</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6 mice</td>
			<td>Xipu&#8217;er-bikai Experimental Animal Co., Ltd (Shanghai, China)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hair removal cream</td>
			<td>Veet, France</td>
			<td>VT-200</td>
			<td></td>
		</tr>
		<tr>
			<td>Hematoxylin and eosin Kit&#160;</td>
			<td>Beyotime biotech&#160;</td>
			<td>C0105M&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>RWD Life Science,China</td>
			<td>R510-22-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>&#160;Yuyan Instruments, China</td>
			<td>SM-301</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical suture needles</td>
			<td>NINGBO MEDICAL NEEDLE CO.,LTD, China</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Thermostatic heating platform</td>
			<td>Qingdao Juchuang Environmental Protection Group Co., Ltd, China</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasound device</td>
			<td>FUJIFILM VisualSonics, Inc.</td>
			<td>Vevo 2100</td>
			<td>Image modes includes B-Mode, Color Doppler Mode and Pulsed Wave Doppler Mode</td>
		</tr>
		<tr>
			<td>Ultrasound gel</td>
			<td>Parker Laboratories,United States</td>
			<td>REF 01-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasound transducer</td>
			<td>FUJIFILM VisualSonics, Inc.</td>
			<td>MS 400</td>
			<td></td>
		</tr>
	</tbody>
</table>

establishment and confirmation of a postnatal right ventricular volume overload mouse model

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.

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

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

Our research scope is the congenital heart disease and heart regeneration. We are trying to understand how volume or pressure overload affects the postnatal heart and lung development, including cardiomyocyte proliferation and maturation. We have developed various types of postnatal mouse or rat models of congenital heart disease.For example, ventricular overload, pressure overload, and the reduced pulmonary blood flow. These models are fascinating and would advance our understanding of pediatric congenital heart disease. For decades, the AVF model has been used to induce chronic heart VO in adult animals.However, little is known about the model in postnatal mice. This study generated a VO postnatal mouse model by creating an abdominal AVF. The morphological and the hemodynamic changes of the right ventricle are demonstrated.

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 po...

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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 ...

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699 Views.  Shanghai Jiaotong University. Our research scope is the congenital heart disease and heart regeneration. We are trying to understand how volume or pressure overload affects the postnatal heart and lung development, including cardiomyocyte proliferation and maturation. We have developed various types of postnatal mouse or rat models of congenital heart disease.For example, ventricular overload, pressure overload, and the reduced pulmonary blood flow. These models are fascinating and would advance our understanding o...

Video: Author Spotlight: Establishment and Confirmation of a Postnatal Right Ventricular Volume Overload Mouse Model  

Abdominal Arteriovenous Fistula Surgery in Postnatal Mice to Establish RV Volume Overload Model

Begin by testing for pinch reflexes on the anesthetized mouse pup's tail to confirm the depth of anesthesia. Next, disinfect the skin with three alternating rounds of Betadine and 70%ethanol using sterile cotton swabs. Cut the abdominal wall and peritoneum from the lower abdomen to the subxiphoid to fully expose the peritoneal cavity.Use cotton swabs to gently pull away the gastrointestinal tract and the bladder to visualize the vertical abdominal aorta or AA and the inferior vena cava or IVC under the retroperitoneum. After rotating the operating table 90 degrees counterclockwise, adjust the microscope magnification to view the two horizontal vessels clearly. Using an 11-0 suture needle having a 0.7 millimeter diameter, puncture the fistula obliquely from the AA into the IVC.Verify the successful puncture by observing the swelling and mixing of the venous and arterial blood in the IVC. Press the bleeding point with dry cotton swabs for 15 seconds to compress it rapidly. Immediately place the stomach, intestines and bladder back into the abdominal cavity to promote hemostatic compression.Make a blanket stitch on the abdominal wall and peritoneum using a 9-0 suture thread, then place the mouse on a warm pad.

Confirmation of Established Abdominal Arteriovenous Fistula by Ultrasound and Volume Overload by Echocardiography in Postnatal Mice

To confirm the fistula by ultrasound, begin by taping down the anesthetized mouse in a supine position on the warm platform. Then connect the mouse to an ECG monitor with pre-warmed ultrasound gel. Use hair removal cream to depilate the chest and abdominal skin.Place a 24 megahertz transducer on the mid-abdominal line and rotate the transducer marker to the mouse head. Now, move the platform down to the left or right side of the mouse and use B-mode and Color Doppler mode to visualize the long axis view of the vessels and blood signals. Use the Pulsed-Wave Doppler mode to measure the blood flow velocity of the AA, IVC, and fistula to confirm arteriovenous fistula patency.To confirm ventricular volume overload, move the tail end of the platform downward. Then place the transducer on the chest rotating the marker to the right shoulder of the mouse. Use the B-mode and the Color Doppler modes to visualize the modified parasternal long axis view of the pulmonary artery or PA.Now using the Pulsed Wave Doppler mode measure the blood flow signals in the PA, including the velocity, time integral, PA valve diameter, pulmonary arterial acceleration, and right ventricular ejection time.Use the mean of three consecutive measurements in each case. The postnatal VO mice showed an acceptable perioperative survival rate in the early postoperative period. The AVF patency rates were 70%at one and two weeks, 46.7%at one month, and 40%at two months postoperatively.Both the PA valve diameter and the right ventricular stroke volume increased within two months. The PA velocity time integral was relatively higher in the VO group within two weeks postoperatively, but declined thereafter. The PA flow patterns changed with decreasing PA acceleration.The right ventricular systolic pressure significantly increased with pulmonary regurgitation two months after surgery. Microscopic analysis showed a significantly enlarged right ventricle of the VO group after AVF, while histological staining showed a thickened RV3 wall and enlarged RV cavity.