A Neonatal Heterotopic Rat Heart Transplantation Model for the Study of Endothelial-to-Mesenchymal Transition

Summary

This work presents an animal model of endothelial-to-mesenchymal transition-induced fibrosis, as seen in congenital cardiac defects such as critical aortic stenosis or hypoplastic left heart syndrome, which allows for detailed histological tissue evaluation, the identification of regulatory signaling pathways, and the testing of treatment options.

Abstract

Endocardial fibroelastosis (EFE), defined by subendocardial tissue accumulation, has major impacts on the development of the left ventricle (LV) and precludes patients with congenital critical aortic stenosis and hypoplastic left heart syndrome (HLHS) from curative anatomical biventricular surgical repair. Surgical resection is currently the only available therapeutic option, but EFE often recurs, sometimes with an even more infiltrative growth pattern into the adjacent myocardium.

To better understand the underlying mechanisms of EFE and to explore therapeutic strategies, an animal model suitable for preclinical testing was developed. The animal model takes into consideration that EFE is a disease of the immature heart and is associated with flow disturbances, as supported by clinical observations. Thus, the heterotopic heart transplantation of neonatal rat donor hearts is the basis for this model.

A neonatal rat heart is transplanted into an adolescent rat's abdomen and connected to the recipient's infrarenal aorta and inferior vena cava. While perfusion of the coronary arteries preserves the viability of the donor heart, flow stagnation within the LV induces EFE growth in the very immature heart. The underlying mechanism of EFE formation is the transition of endocardial endothelial cells to mesenchymal cells (EndMT), which is a well-described mechanism of early embryonic development of the valves and septa but also the leading cause of fibrosis in heart failure. EFE formation can be macroscopically observed within days after transplantation. Transabdominal echocardiography is used to monitor the graft viability, contractility, and the patency of the anastomoses. Following euthanasia, the EFE tissue is harvested, and it shows the same histopathological characteristics as human EFE tissue from HLHS patients.

This in vivo model allows for studying the mechanisms of EFE development in the heart and testing treatment options to prevent this pathological tissue formation and provides the opportunity for a more generalized examination of EndMT-induced fibrosis.

Introduction

Endocardial fibroelastosis (EFE), defined by the accumulation of collagen and elastic fibers in the subendocardial tissue, presents as a pearly or opaque thickened endocardium; EFE undergoes most active growth during the fetal period and early infancy¹. In an autopsy study, 70% of cases with hypoplastic left heart syndrome (HLHS) were associated with the presence of EFE².

Cells expressing markers for fibroblasts are the main cell population in EFE, but these cells also concomitantly express endocardial endothelial markers, which is an indication of the origin of these EFE cells. Our group previously established that the underlying mechanism of EFE formation involves a phenotypical change of endocardial endothelial cells to fibroblasts through endothelial-to-mesenchymal transition (EndMT)³. EndMT can be detected using immunohistochemical double-staining for endothelial markers such as cluster of differentiation (CD) 31 or vascular endothelial (VE)-cadherin (CD144) and fibroblast markers (e.g., alpha-smooth muscle actin, α-SMA). Furthermore, we also previously established the regulatory role of the TGF-ß pathway in this process with activation of the transcription factors SLUG, SNAIL, and TWIST³.

EndMT is a physiological process that occurs during embryonic cardiac development and leads to the formation of the septa and valves from endocardial cushions⁴, but it also causes organ fibrosis in heart failure, kidney fibrosis, or cancer and plays a key role in vascular atherosclerosis^5,6,7,8. EndMT in cardiac fibrosis is mainly regulated through the TGF-β pathway, as we and others have reported^3,9. Various stimuli have been described to induce EndMT: inflammation¹⁰, hypoxia¹¹, mechanical alterations¹², and flow disturbances, including alterations of the intracavitary blood flow¹³, and EndMT may also be a consequence of a genetic disease¹⁴.

This animal model was developed using the key components of cardiac EFE development, which are immaturity and alterations of the intracavitary blood flow, specifically flow stagnation. Immaturity was fulfilled by using neonatal rat hearts as donors, since neonatal rats are known to be developmentally immature immediately after birth. Heterotopic heart transplantation offered the provision of intracavitary flow restriction¹⁵.

From a clinical point of view, this animal model allows for better investigating the impact of EndMT on the growing left ventricle (LV). The growth restriction imposed on the fetal and neonatal heart through EndMT-induced EFE formation¹⁶ precludes patients with left ventricular outflow tract obstructions (LVOTO) such as congenital critical aortic stenosis and hypoplastic left heart syndrome (HLHS) from curative anatomical biventricular surgical repair¹⁷. This animal model facilitates the study of the cellular mechanisms and regulation of tissue formation through EndMT and allows for the testing of pharmacological treatment options^3,18.

Transabdominal echocardiography is used to monitor the graft viability, contractility, and the patency of the anastomoses. Following euthanasia, EFE formation can be macroscopically observed within 3 days after transplantation. EFE tissue shows the same histopathological characteristics as human EFE tissue from patients with LVOTO.

Hence, this animal model, though developed for pediatric use in the spectrum of HLHS, can be applied when studying various diseases based on the molecular mechanism of EndMT.

Protocol

All the animal procedures were conducted in accordance with the National Research Council. 2011. Guide for the Care and Use of Laboratory Animals: Eighth Edition. The animal protocols were reviewed and approved by the Institutional Animal Care and Use Committee at Boston Children's Hospital.

Prior to surgery, all the surgical instruments are steam-autoclaved, and modified Krebs-Henseleit buffer, with a final concentration of 22 mmol/L KCl, is prepared as a cardioplegic solution (Table 1). The solution is filter-sterilized and stored at 4 °C overnight. A surgical microscope (12.5x) is required for the heterotopic neonatal rat heart transplantation procedure.

1. Preparation and anesthesia

1. Use male/female Lewis rats with a weight of around 150 g (5-6 weeks of age) as recipients.
2. To start, generously shave the rat's abdomen with a razor.
3. Place the rat into an isoflurane chamber, and turn on the oxygen flow at 2 L/min with 2% isoflurane until the animal is properly sedated but still spontaneously breathing. Inject 45 mg/kg ketamine and 5 mg/kg xylazine intraperitoneally (IP), as well as 300 U/kg heparin. Confirm proper anesthetization with a toe pinch test.
   NOTE: Carefully monitor the spontaneous breathing and heart rate through palpation of the chest to assure a stable hemodynamic status throughout the entire process.
4. For intubation, place the rat on an oblique shelf (Figure 1), secure the front teeth with a string, and place the head facing toward the surgeon.
5. Place the light on the outside of the neck onto the area of the vocal cords, grab the tongue with two fingers, and slightly push it upward and to the left to provide optimal vision for intubation. Use an 18 G, 2 in cannula for a 100-150 g rat. Secure the intratracheal tube with tape.
   NOTE: Surgical loups with 3.5x magnification are recommended for intubation.
6. Connect the intubation cannula to the small animal ventilator, and adjust the settings according to the manufacturer's instructions based on the animal size.
   NOTE: Use the following settings for a 150 g rat: volume mode; respiratory rate, 55/min; tidal volume, 1.3 mL 50 % I/E ratio, but this can be adjusted appropriately as needed. Assure proper bilateral and equal chest movement, and administer isoflurane continuously at 0.5%–2% through the ventilator.
7. Place the rat on a heating pad (to maintain normal body temperature) in a supine position with the tail facing toward the surgeon. Sterilize the abdomen three times with betadine solution and 70% of ethanol alternatingly. Administer eye lube, and cover the rat with a sterile surgical drape, leaving the abdomen uncovered.

2. Surgical preparation and heterotopic transplantation of the neonatal donor heart in the recipient rat

1. Perform a midline laparotomy using a 15 blade scalpel for the skin incision, and use scissors to open the anterior abdominal wall, followed by blunt exposure of the retroperitoneal abdominal aorta and inferior vena cava (IVC) with cotton tip applicators.
2. Mobilize the intestines (including the descending colon), and place them toward the right upper quadrant. Cover the intestines with warm saline-soaked gauze. Use retractors to ensure optimal exposure of the IVC and abdominal aorta.
3. Perform blunt dissection of the infrarenal IVC and abdominal aorta up toward the bifurcation. Ligate all the infrarenal branching arteries and veins (e.g., inferior mesenteric artery and lymph node arteries) with a 10-0 nylon suture.
   NOTE: There is great variability in the anatomy of these side branches. Monitor the aorta's pulse and heart rate visually when no other hemodynamic monitoring is available. Assess the proper depth of anesthesia every 15 min through a toe pinch test. Adjust the isoflurane concentration accordingly.
4. After the donor heart is harvested from a neonatal rat, deliver the excised heart in sterile conditions in a surgical basin containing Krebs-Henseleit buffer to the surgical field. Irrigate the donor heart intermittently with ice-cold cardioplegic solution.
   NOTE: When a second surgeon is available, the heart should be prepared at the same time, as a second surgeon reduces the total anesthesia time of the recipient animal and the ischemia time of the donor heart. When a second surgeon is not available, cover the recipient's abdomen with warm saline, and monitor the animal during the harvesting procedure.
5. Apply four small atraumatic vascular clamps to the distal and proximal segments of the infrarenal aorta and IVC. If needed, temporarily occlude an unfavorable renal vessel with a 7-0 silk suture, and release the suture after the procedure. Place a 10-0 nylon suture vertically onto the anterior wall of the aorta to facilitate the aortotomy. Perform an aortotomy with two small horizontal cuts (wedge-shaped) with microscissors by slightly pulling up the suture.
   NOTE: To remove any blood clots, flushing of the aortic lumen with heparinized saline is recommended.
6. Place the donor heart on the left side (from the animal’s perspective) of the aorta and secure the recipient’s infrarenal aorta and the donor’s ascending aorta end-to-side at the 12 o’clock and 6 o’clock positions of the aortotomy with sutures. Continue with the third and fourth sutures at the 3 o’clock and 9 o’clock positions, gently flipping the heart over to the right side of the aorta after the third suture. Complete the arterial anastomosis by adding one to two sutures to every interspace.
   NOTE: Care should be taken to avoid touching either the donor's ascending aorta or the recipient's abdominal aorta with forceps when creating the anastomosis to avoid tissue damage.
7. Rotate the rat counterclockwise, with the head facing toward the surgeon's left hand. Move the donor's aorta to the left side of the abdominal aorta to allow optimal sight onto the IVC.
8. Perform a venotomy on the IVC, slightly proximal to the aortic anastomosis, using an 11 blade for puncture and microscissors for adequate size adjustment according to the diameter of the donor's pulmonary trunk. Again, flush the intracaval lumen with heparinized saline.
9. Start with the venous anastomosis between the recipient's IVC and donor's pulmonary trunk, which is best achieved by placing interrupted 11-0 nylon sutures on the back wall of the vessel, starting at the 12 o'clock and 6 o'clock positions (related to the IVC), and then place a continuous 11-0 nylon suture on the front wall (from the 6 o'clock toward the 12 o'clock position).
10. Cover the anastomoses with small strips of an absorbable gelatin sponge, and remove the microvascular clamps starting distally. Use a cotton tip applicator to lightly compress the sponges to obtain optimal hemostasis.
11. Observe the graft's coronary vessels filling at the time of the release of the distal microvascular clamps, and make sure that the donor heart starts beating immediately when the proximal clamp is released.
    NOTE: The graft's viability can be scored from 0 to 4 intraoperatively according to a modified Stanford score¹⁹ to confirm adequate graft function.
12. Place the intestines back into the abdomen by ensuring not to distort the arterial and venous anastomosis.
13. Administer meloxicam (1mg/kg) and ethiqa XR (0.65 mg/kg) subcutaneously while the animal is fully anesthezised to ascertain postoperative analgesia. Then, close the abdominal wall with a continuous 5-0 absorbable vicryl suture before closing the skin with a 6-0 absorbable vicryl suture intracutaneously.
    ​NOTE: Guidance regarding common failures and troubleshooting is presented in Table 2.

3. Harvesting of the neonatal donor heart

1. Place the neonatal donor rat in a chamber insufflated with isoflurane (2%) for sedation. Administer ketamine (75 mg/kg) and xylazine (5 mg/kg), as well as heparin (300 U/kg) intraperitoneally.
2. Confirm the depth of anesthesia by toe pinch, and place the rat in a supine position with the tail facing toward you. Sterilize the entire thorax and abdominal wall with betadine and 70% ethanol three times alternatively. Cover the rat with a sterile surgical drape.
3. Using a 12.5x surgical microscope, remove the entire anterior thoracic wall by starting with a horizontal incision using a 15 blade scalpel at the xyphoid followed by vertical incisions laterally up to the axillae on both sides with scissors. The anterior thoracic wall can then be removed by continuing with another horizontal incision right beneath the neck.
4. Dissect the IVC, right and left superior vena cavae, and pulmonary vessels with scissors, and then encircle and ligate all the vessels with a 7-0 silk suture. Administer 3 mL of ice-cold, high-potassium modified Krebs-Henseleit solution to the right atrium by puncturing the IVC with a 30 G needle and slightly pushing the diaphragm down with forceps.
5. Cut the IVC, SVCs, pulmonary vessels, and aorta with scissors. Transect the pulmonary arteries as far as possible and the aorta distal to the brachiocephalic trunk to ensure proper length using a 11 blade scalpel.
6. Separate the pulmonary trunk and ascending aorta with microscissors, and flush the heart with ice cold cardioplegic solution using a 3 mL syringe.

4. Recovery of the recipient and graft monitoring

1. After surgery, give the rat ample time to wake up, which usually occurs in a 15 min time window, and let it recover on a heating pad.
   NOTE: No antibiotics are necessary due to the very low risk of infection and in order to not compromise the experimental model, and no restriction to food or water is applied.
2. Following transplantation, monitor the graft function by palpation of the transplanted heart daily, but consider that this can sometimes be difficult to assess due to intestine overlay.
   NOTE: Abdominal echocardiography can more accurately measure the graft viability. For echocardiography, sedate the rat slightly with isoflurane (1-2%) inhaled through a nose cone, and position it on a heating pad. Echocardiography is usually performed on postoperative day (POD) 1, POD 7, and POD 14. To allow for the assessment of the heart rate and contractility, one can easily obtain long-axis and short-axis views (Figure 2A, B). To evaluate the anastomoses, use Doppler echocardiography (Figure 3A), and confirm the formation of EFE tissue as seen as an echo-bright endocardial layer within the left ventricular cavity (Figure 3B, C).

Results

Graft viability and beating
In this work, the graft viability was visually assessed after all the clamps had been removed, and an approximate reperfusion time of 10-15 min was allowed with an open abdomen for observation of the graft. The same scoring system to objectively verify graft viability was used for visual assessment at the end of surgery and for the echocardiography on POD 1, POD 7, and POD 14.

0 = no organ function; 1 = (rest) organ function, only minimal contraction; 2 = weak or partial organ function; 3 = contractile rate or intensity reduced, but homogenous organ function; 4 = optimal atrium and ventricle contraction (120-160 beats/min). A score of 3 or 4 was rated a success. Palpatory evaluation of the abdominal donor graft was used to monitor the graft viability between the time points of echocardiographic assessment.

Mortality and graft viability success rate
The procedure was introduced to a new surgical team at the study center between October 2022 and December 2022, and 19 neonatal heterotopic rat heart transplantations were performed at the study center during this period. The immediate operative survival rate was 79%, and the graft viability success rate (displaying a viable, beating donor heart) was 84%. The procedure characteristics are presented in Table 3.

Among the 12 surviving animals, 2 required euthanasia prior to the 2 week study endpoint, 1 due to an ileus (n = 1), and the other due to pain unrelieved with pain medication (n = 1), and 2 were euthanized by design 1 week after surgery.

In three rats, the applied modified Stanford score increased from 3 to 4 between immediate postoperative visual grading and echocardiographic evaluation on POD 1. Among the eight surviving rats at the 14 days endpoint, the modified Stanford scores at echocardiography were four for seven animals and three for one animal. The most common cause of death in this series was hemodynamic failure due to excessive blood loss as a consequence of the very immature heart and, thus, fragile donor vessels for anastomosis or long anesthesia times.

Histological assessment of EFE tissue
Following CO₂ euthanasia of the recipient rat, a re-laparotomy was performed under sterile preparation. The donor graft was excised and immediately placed in a physiologic saline solution on ice for further processing. A horizontal slice was resected at the mid-ventricular level of the right and left ventricle, placed in optimal cutting temperature (OCT) embedding medium, and frozen in liquid nitrogen (Figure 4A). All the other tissue was snap-frozen with liquid nitrogen and stored in a −80°C freezer for further analysis. Images were acquired using an inverted microscope (Figure 4B-D).

Immunohistochemical staining as the gold standard to identify EndMT was performed using 4',6-diamidino-2-phenylindole (DAPI) (blue), VE-Cadherin as an endothelial marker (red), and α-SMA as a fibroblast marker (green). Phosphorylated SMAD proteins and the transcription factor SLUG/SNAIL were also stained in the EFE tissue (Figure 5A-E)^3,20.

Figure 1: Oblique shelf for intubation. The rat is placed on its back, with the front teeth secured with a string and the head facing toward the surgeon. Please click here to view a larger version of this figure.

Figure 2: Echocardiographic long-axis view of the LV. (A) Native rat heart indicating normal filling during diastole. (B) Donor graft with flow stagnation within the LV. Diminished volume loading during diastole. Abbreviations: LV = left ventricle; MV = mitral valve; LVOT = left ventricular outflow tract. Please click here to view a larger version of this figure.

Figure 3: Anastomoses and EFE evaluation. (A) Echocardiographic color Doppler study indicating patent arterial (red arrow) and venous (blue arrow) anastomoses. (B,C): Echo-bright endocardial surface within the LV cavity indicative of EFE (white arrows). Abbreviations: LV = left ventricle; EFE = endocardial fibroelastosis. Please click here to view a larger version of this figure.

Figure 4: Macroscopic and microscopic tissue evaluation. (A) Mid-ventricular cross-section through the LV and RV. The white arrows point toward the EFE tissue. (B) Hematoxylin-eosin, (C) Masson's trichrome (MTS), and (D) Elastin van Gieson (EVG) staining. The large magnification indicates that the EFE tissue (black arrows) contains high amounts of organized collagen (blue in MTS) and elastin fibers (black in EVG). Abbreviations: LV = left ventricle; RV = right ventricle; EFE = endocardial fibroelastosis. Please click here to view a larger version of this figure.

Figure 5: Comparison of histological and immunohistological images. (A) Hematoxylin-eosin staining. (B-E) Immunohistochemical staining; EFE tissue double-stained for (B,C) VE-Cadherin and α-SMA, (D) CD31 and phospho-SMAD2/SMAD3 (colocalized with the nuclei stained with DAPI in blue), and (E) CD31 and SLUG/SNAIL (colocalized with the nuclei stained with DAPI in blue), indicative of EndMT, as shown by the white arrows. Abbreviations: LV = left ventricle; EFE = endocardial fibroelastosis; EndMT = endothelial-to-mesenchymal transition. Please click here to view a larger version of this figure.

1 liter of sterilized, distilled water
NaCl	118 mmol/L
KCl	22 mmol/L
KH2PO4	1.2 mmol/l
MgSO4	1.2 mmol/L
NaHCO3	25 mmol/L
Glucose	11 mmol/L
CaCl2	2.5 mmol/L

Table 1: Composition of the Modified Krebs-Henseleit buffer. High-potassium (22 mmol/L KCl) cardioplegic solution is prepared, filter-sterilized, and stored at 4 °C overnight.

Common failures and troubleshooting
Graft does not start to beat/coronary arteries do not fill after releasing the clamps	Check for thrombus formation at arterial anastomosis
	Check for ischemia time (=total arrest time) (should not exceed 100 minutes)
Long wake-up time or rat does not wake up after surgery	Monitor pulse strength and frequency during surgery and reduce isoflurane inhalation, if hemodynamics are weak
	Immediate postoperative livid or necrotic intestines are suspicious for reduced intraoperative hemodynamics often due to long anesthesia time
Weak hemodynamics right after laparotomy	Adjust isoflurane flow for anesthesia
	Evaluate intubation and proper chest motion: unilateral intubation, pneumothorax, obstructed endotracheal lumen are common failures in the beginning.
Rat wakes up but dies in first 24 hours	Extensive blood loss during surgery
	If increased amount of blood is found at autopsy in abdomen, it is most likely due to failure of the anastomosis

Table 2: Common failures and troubleshooting. Thorough monitoring and re-evaluation of unsuccessful procedures are crucial to achieving a high survival rate in this model.

Recipient rat weight in grams, median [IQR]	150 [50]
Donor age in days, median [IQR]	3 [1]
Donor weight in grams, median [IQR]	9 [2]
Graft ischemia time in minutes, median [IQR]	100 [25]
Postoperative success rate, n	16/19 (=84%)

Table 3: Procedure characteristics. Recipient and donor selection, graft ischemia time, and survival rate. Abbreviation: IQR = interquartile range.

Discussion

This animal model of heterotopic transplantation of a neonatal donor rat heart into the recipient's abdomen creates the possibility to study EndMT-derived fibrosis through detailed histological tissue evaluation, identify regulatory signaling pathways, and test treatment options. Since EndMT is the underlying mechanism for fibrotic diseases of the heart, this model has great value in the field of pediatric cardiac surgery and beyond. In this model, many factors can negatively influence the outcome of the procedure. Thus, proper handling of the very fragile tissue due to the immaturity of the donor heart, proper animal handling during anesthesia, and high-level microsurgical skills are basic requirements for the success of this model. An optimal technical setup, including a surgical microscope, small animal ventilator, and microsurgical instruments, should be used when performing these experiments. Although not essential, basic monitoring of the heart rate or body temperature could be beneficial, especially for inexperienced surgeons, in order to monitor the hemodynamics and depth of anesthesia.

Important surgical aspects to bear in mind include the immaturity of the neonatal donor hearts, which makes the tissue very fragile and leaves the ascending aorta and pulmonary trunk vulnerable to tears. Thus, any handling should be undertaken with great care. Due to the small vessels used for anastomosis, it is recommended to perform arterial anastomosis with interrupted stitches and intermittent flushing of the anastomosis site with heparinized saline, which helps to avoid thrombus formation. Selection of appropriately aged neonatal rats is required to overcome the issue of using hearts that are too immature and, therefore, highly susceptible to anastomosis rupture. On the other hand, after a certain age of about 7 days, EndMT can no longer be shown reproducibly in this animal model¹⁵.

EndMT has been identified as the central mechanism for various kinds of cardiac fibrosis and atherosclerosis, but research has been hampered due to a lack of in vivo models⁸. The main developments in the field of EndMT research are restricted to cell culture models, which have inherent limitations^3,8,9. Furthermore, studies on endocardial endothelial cells are even more restricted. As an alternative, coronary artery endothelial cells are often used as a substitute, as they have been reported to originate in part from endocardial cells²¹. Hence, this animal model can be used not only for cardiac fibrosis but to study important pathomechanisms of flow-induced EndMT in atherosclerosis. For congenital heart disease, we have shown the ability to reproduce the transition from healthy endocardium to EFE tissue through EndMT in our rat model, with EFE that structurally resembles human EFE tissue. There is some controversy regarding the cellular origin of mesenchymal cells within the EFE tissue. Clark et al.²² reported that epicardial cells contribute to EFE, but our data indicated that the majority of EFE tissue is derived through endocardial endothelial cells undergoing EndMT³. Experiments on a single-cell level are currently underway to ascertain the cellular origin of EFE tissue.

Through this in vivo model, the regulatory pathways of EndMT can be studied. An imbalance, specifically an increase in the TGF-ß pathway and impaired bone morphogenetic protein (BMP) signaling, has been shown to play a major role in endocardial cells expressing transcription factors regulating EndMT. Alternatively, Jagged/NOTCH signaling and Wnt/ß-Catenin have also been reported to induce EndMT^3,23. The TGF-ß pathway induces the activation of transcription factors like SLUG, SNAIL, and TWIST via SMAD proteins, thereby regulating EndMT^20,24. In this animal model, we have been able to recapitulate these mechanisms, which have been confirmed by immunohistochemical staining.

The stimulating factors for EndMT-induced fibrosis in this animal model are immaturity and flow stagnation, whereas other models are designed to induce EndMT through genetic modifications, hypertension, or dietary restrictions^9,25. Compared to other species, neonatal rats are very immature at birth, and therefore, they are particularly susceptible to undergoing EndMT.

We and others have used mice to better study the origins of EFE via transgenic lineage tracing, but several limitations need to be discussed^3,22. First, due to the model's complexity, mortality rates are higher in mice compared to rats, and the presentation of EFE is more heterogenous; therefore, the rat model is more reliable and reproducible. Echocardiographic measures are crucial to assess the graft function throughout the study period, and we have shown that with these measures, as well as assessing the pulsatility and patency of the anastomoses, graft function, and contractility can also be studied. With more experience, even more advanced analyses of the transplanted heart, such as strain analysis of the LV, could be performed in rat models. It is currently unclear whether the same pathophysiological condition can be induced in larger animals other than rodents, and this requires further investigation.

In conclusion, this pediatric animal model mimics the human disease of EndMT and can be useful to determine the regulation of EndMT and to study pharmacological interventions to inhibit this pathological process.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Advanced Ventilator System For Rodents, SAR-1000	CWE, Inc.	12-03100	small animal ventilator
aSMA	Sigma	A2547	Antibody for Immunohistochemistry
Axio observer Z1	Carl Zeiss		inverted microscope
Betadine Solution	Avrio Health L.P.	367618150092
CD31	Invitrogen	MA1-80069	Antibody for Immunohistochemistry
DAPI	Invitrogen	D1306	Antibody for Immunohistochemistry
DemeLON Nylon black 10-0	DemeTECH	NL76100065F0P	10-0 Nylon suture
ETFE IV Catheter, 18G x 2	TERUMO SURFLO	SR-OX1851CA	intubation cannula
Micro Clip 8mm	Roboz Surgical Instrument Co.	RS-6471	microvascular clamps
Nylon black monofilament 11-0	SURGICAL SPECIALTIES CORP	AA0130	11-0 Nylon
O.C.T. Compound	Tissue-Tek	4583	Embedding medium for frozen tissue specimen
p-SMAD2/3	Invitrogen	PA5-110155	Antibody for Immunohistochemistry
Rodent, Tilting WorkStand	Hallowell EMC.	000A3467	oblique shelf for intubation
Silk Sutures, Non-absorbable, 7-0	Braintree Scientific	NC9201231	Silk suture
Slug/Snail	Abcam	ab180714	Antibody for Immunohistochemistry
Undyed Coated Vicryl 5-0 P-3 18"	Ethicon	J493G	5-0 Vicryl
Undyed Coated Vicryl 6-0 P-3 18"	Ethicon	J492G	6-0 Vicryl
VE-Cadherin	Abcam	ab231227	Antibody for Immunohistochemistry
Zeiss OPMI 6-SFR	Zeiss		Surgical microscope
Zen, Blue Edition, 3.6	Zen		inverted microscope software