Endocardial fibroelastosis hinders left ventricular growth and prevents biventricular surgical repair for patients with congenital critical aortic stenosis and hypoplastic left heart syndrome. While surgical resection is currently the only available treatment option, EFE frequently recurs. Therefore, we aim to investigate the mechanisms behind EFE and develop new therapeutic strategies using an animal model.
The main developments in the field of NDMT research are restricted to cell culture models. Which have inherent limitations, and research has been hampered due to a lack of in vivo models. Our group previously established that the underlying mechanism of EFE formation involves a phenotypical change of endocardial endothelial cells to fibroblasts.
A process called endothelial-to-mesenchymal transition. This animal model allows for a more generalized examination of NDMT-induced fibrosis in the heart. Whereas other models are designed to induce NDMT through genetic modifications, hypertension, or dietary restrictions.
As EFE often recurs after primary resection, sometimes with an even more infiltrative growth pattern, we are especially interested in the molecular interaction with the underlying myocardium. Begin harvesting the neonatal donor heart by anesthetizing and sterilizing the rat. And making a horizontal incision at the xiphoid with a 15 blade scalpel under a 12.5X surgical microscope.
Next, use scissors to create vertical incisions laterally up to the axillae on both sides. Remove the anterior thoracic wall with another horizontal incision beneath the neck. Dissect the inferior vena cava, or IVC, right and left superior vena cavi, or SVC, and pulmonary vessels with scissors.
Encircle and ligate all vessels using a 7-0 silk suture. Slightly push the diaphragm down with forceps. And administer three milliliters of ice cold high potassium modified Krebs-Henseleit solution to the right atrium by puncturing the IVC with a 30 gauge needle.
Cut the IVC, SVC, and pulmonary vessels using scissors. Ensure proper length by transecting the pulmonary arteries as far as possible. And the aorta distal to the brachiocephalic trunk using a number 11 blade scalpel.
Flush the heart with the ice cold cardioplegic solution using a three milliliter syringe. Place the anesthetized rat on an oblique shelf. Secure its front teeth with a string, and position the head towards the surgeon.
Place the light on the outside of the neck onto the area of the vocal cords. Grasp the tongue with two fingers, and gently push it upwards and to the left for optimal intubation visibility. Now use an 18 gauge two inch cannula for a 100 to 150 gram rat, and secure the intratracheal tube with tape.
Connect the intubation cannula to the small animal ventilator. And adjust the settings based on the animal size, following the manufacturer's instructions. After intubating the rat, perform a midline laparotomy using a 15 blade scalpel to make the skin incision.
Open the anterior abdominal wall with scissors. And mobilize the intestines toward the right upper quadrant. Next, expose the retroperitoneal abdominal aorta and IVC using cotton tip applicators.
Cover the intestines with warm saline-soaked gauze. And use retractors to maintain optimal exposure of the IVC and abdominal aorta. Now perform blunt dissection of the infrarenal IVC and abdominal aorta up to the bifurcation.
Ligate all infrarenal branching arteries and veins with a 10-0 nylon suture. Transport the harvested donor's heart in a sterile surgical basin containing krebs-henseleit buffer to the surgical field. Ensure to irrigate the donor heart intermittently with ice cold cardioplegic solution.
Next, separate the pulmonary trunk and descending aorta using micro scissors. And irrigate the donor heart with ice cold cardioplegic solution. Next, apply four atraumatic vascular clamps to the distal and proximal segments of the infrarenal aorta and IVC.
Perform an aortotomy using micro scissors to make two small horizontal cuts. Then, place the donor heart on the left side of the aorta. Secure the recipient's infrarenal aorta and the donor's ascending aorta end to side at the 12 and six o'clock positions of the aortotomy with sutures.
Proceed with the third and fourth sutures at the three and nine o'clock positions. Gently flipping the heart to the left side of the aorta after the third suture. Finalize the arterial anastomosis by adding one to two sutures to each interspace.
Now, rotate the rat counterclockwise so the head faces the surgeon's left hand. Shift the donor's heart to the left side of the abdominal aorta for an optimal view of the IVC. Perform a venotomy on the IVC slightly proximal to the aortic anastomosis.
Using an 11 blade for puncture, and micro scissors to adjust the size according to the donor's pulmonary trunk diameter. Flush the intracaval lumen with heparinised saline. Next, perform the venous anastomosis between the recipient's IVC and the donor's pulmonary trunk by placing interrupted 11-0 nylon sutures on the back wall of the vessel, starting at the 12 and six o'clock positions.
Place a continuous 11-0 nylon suture on the front wall. Place absorbable gelatin sponge strips over the anastomosis. And remove the microvascular clamps beginning with the distal ones.
Ensure that the coronary vessels fill. And that the donor's heart begins beating shortly upon release of the proximal clamps. Apply light pressure using a cotton tip applicator on the sponges to achieve optimal hemostasis.
Carefully place the intestines back into the abdomen, being cautious not to distort the arterial and venous anastomosis. Close the abdominal wall and allow the animal to recover. A horizontal slice of the donor graft resected at the mid ventricular level of the right and left ventricles is shown.
Histological analysis indicated that the endocardial fibroelastosis tissue had high amounts of organized collagen and elastin fibers. Endocardial fibroelastosis tissue double stained for VE-Cadherin and Alpha-SMA. CD-31, phospho-SMAD2, and SMAD3.
CD-31 and SLUG or SNAIL indicated endothelial-to-mesenchymal transition.
