This work was supported by a grant B2021-0991 from the Chonnam National University Hospital Biomedical Research Institute and NRF-2020R1F1A1073921 from the National Research Foundation of Korea

The ethical committee of the Laboratory Animal Research Center of Chonnam National University Hospital (approval no. CNU IACUC - H - 2022-36) approved all the animal experiments. Male Sprague-Dawley rats (350-450 g), used in this study received care in compliance with the guidelines for the care and use of the laboratory animals. The rats were housed in temperature-controlled rooms with a 12 h light-dark cycle, with standard food and water available.
1. Preparation
NOTE: A single experimenter can conduct all experimental procedures.
<ol>
	<li>Assemble the Langendorff apparatus, including the oxygenator, pump, and perfusion lines, prior to surgery (Figure 2). Fill the perfusion circuit with 20 mL of saline solution and circulate it until it is primed with autologous blood. 
	NOTE: The objective of this step is to warm the extracorporeal circuit.</li>
	<li>Attach the cardioplegic line to the circuit via the stopcock attached to the aortic cannula and prepare the syringe pump for the final cardioplegic infusion. 
	NOTE: Ensure the removal of any air bubbles from the perfusion circuit and the cardioplegic line.</li>
	<li>Place the temperature sensor within the reservoir where the donor heart will be stored, maintaining the circuit&#39;s temperature at 37 &#176;C.</li>
	<li>Surgical preparations
	<ol>
		<li>Prepare a separate set of sterile micro-instruments and materials for each donor and recipient rat.
		<ol>
			<li>Prepare the surgical set for the donor: pair of surgical scissors, pair of micro forceps, sharp mosquito forceps, 5-0 silk sutures, cotton swabs, 50 mL syringe, perfusion line for the cardioplegic solution (CPS), syringe pump, 18 G angiocatheter, one set of 5 Fr. femoral catheters, and sterile gauzes.</li>
			<li>Prepare the surgical set for the recipient: microsurgical scissors, wound retractor, pair of micro forceps, mosquito forceps, vascular micro clamps, 1 mL syringe, one 5-0 and 9-0 polypropylene sutures, 5-0 silk sutures, cotton swabs, and sterile gauzes.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Donor heart preservation and blood collection
<ol>
	<li>Induce anesthesia in the donor rat with isoflurane (5%) in the anesthesia chamber and record the rat&#39;s weight before placing it on the surgical table.</li>
	<li>Place the rat in the supine position on the surgical table and administer continuous anesthesia by delivering 2%-2.5% isoflurane with 90% oxygen through a nosecone.</li>
	<li>Verify the depth of anesthesia by checking the lack of response to the toe pinch and the breath frequency, which should be between 50-60 per minute. 
	NOTE: An adequate level of anesthesia is crucial to avoid unnecessary stress and pain to the donor rat.</li>
	<li>Apply eye lubricant and shave the region pubis to the clavicula, where the surgery will be performed. Clean the area with an iodine-based scrub and 70% alcohol.</li>
	<li>Catheterization
	<ol>
		<li>Make a 7 cm midline abdominal incision and bilateral incisions measuring 3 cm from the xiphoid process to the mid-clavicle. Remove the pelt from the thoracic region.</li>
		<li>Using cotton swabs, mobilize the abdominal organs to the left side of the abdomen. Isolate the abdominal aorta from the retroperitoneal fascia and adipose tissues.</li>
		<li>Inject 1,000 IU heparin dissolved in 0.3 mL of isotonic saline through the inferior vena cava (IVC) using a 1 mL syringe. Stop any bleeding from the needle hole by gently compressing with a cotton swab. 
		NOTE: Be cautious of air embolism during injection, as it can lead to cardiac arrest.</li>
		<li>Insert a 5 Fr. femoral catheter into the abdominal aorta (Abd. A). Ensure that the catheter tip reaches the aortic arch. Confirm the catheter location by assessing the approximate length of the inserted part of the catheter.</li>
	</ol>
	</li>
	<li>Blood collection
	<ol>
		<li>Collect around 10 mL of blood via the catheter inserted in the Abd. A.</li>
		<li>Later, dilute the priming blood with isotonic saline until the total volume reaches 12 mL. Add 5 mg of cefazolin dissolved in 0.3 mL of saline and insulin (20 IU).</li>
	</ol>
	</li>
	<li>Cardiac arrest
	<ol>
		<li>Connect the previously prepared CPS perfusion line to the abdominal catheter and start the CPS administration with the syringe pump at a rate of 800 mL/h.</li>
		<li>Open the thoracic cavity from the diaphragm and cut the IVC close to the diaphragm to prevent ventricular distention. Cut the ribs bilaterally along the thoracic spine up to the thoracic inlet. Reflect the mobilized ventral chest wall superiorly with mosquito forceps.</li>
		<li>Remove the thymus entirely using micro forceps to visualize the aortic arch. Apply light compression if thymic arteries bleed.</li>
	</ol>
	</li>
	<li>Extraction
	<ol>
		<li>After administering all the CPS, isolate the aortic arch from the surrounding tissues. Carefully dissect just below the left subclavian artery.</li>
		<li>Transect the brachiocephalic and left common carotid arteries at a distant position, leaving the longer stumps of the aortic arch for easy handling during aorta cannulation. Transect the main pulmonary artery (MPA) as close as possible to the bifurcation. Be cautious not to damage the left atrial appendage.</li>
		<li>Carefully ligate the superior vena cava (SVC) and IVC with 5-0 silk sutures, preventing the obstruction of the right atrium (RA) and coronary sinus. Cover the left margins of the thorax with wet gauze, place the heart, onto it and gently retract the SVC and IVC ligatures to expose the hilum.</li>
		<li>Ligate the pulmonary and azygos veins together with a 5-0 silk suture. Sever the tissue dorsal to the ligature and extract the heart. Examine the heart for any injury. Finally, weigh the heart before aortic cannulation.</li>
	</ol>
	</li>
</ol>
3. Ex situ perfusion
<ol>
	<li>Aorta cannulation and perfusion
	<ol>
		<li>Before aorta cannulation, replace the saline-primed circuit with blood priming.</li>
		<li>Insert the aortic cannula into the aortic arch and secure it with a temporary micro clamp. Ensure that the tip of the cannula is positioned at the brachiocephalic junction.</li>
		<li>Confirm the correct position of the cannula by gently grasping the aorta with micro forceps.</li>
		<li>Start the perfusion at a flow rate of 2-3 mL/min, allowing perfusate to leak from the cannulation site to remove any air bubbles.</li>
		<li>Monitor the perfusion pressure and temperature through the sensor connected to the monitoring system.</li>
		<li>Gently massage the heart with the first and index fingers until venous blood leaks from the main pulmonary artery (MPA).</li>
		<li>Secure the aorta with a 1-0 silk ligature and remove the clamp after verifying all the settings (perfusion circuit, perfusion pressure, temperature).</li>
		<li>Once the permanent ligature is placed, ensure the heart begins to contract within a few seconds and reaches normal rhythm in 60 s. A mean perfusion pressure of 55-65 mmHg with a coronary flow rate of 3-4 mL at 37 &#176;C indicates adequate perfusion.</li>
		<li>Collect 0.15 mL of blood from the reservoir and check the blood gas analysis (BGA) at the beginning of perfusion and every 20 min thereafter. Monitor and record the pH, pCO2, pO2, glucose, hematocrit, potassium, and lactate during perfusion. After 120 min of perfusion, administer 3 mL of Custodiol through the syringe pump at a rate of 250 mL/h to arrest the heart.</li>
	</ol>
	</li>
</ol>
4. Implantation
<ol>
	<li>Preparation of recipient
	<ol>
		<li>Begin the recipient preparation 30 min before the cessation of ex situ perfusion.</li>
		<li>Anesthetize the recipient animal using the same method as mentioned in step 2.2.</li>
		<li>Place the rat in a supine position on the heating pad and insert the temperature probe into the rectum to maintain the body temperature at 37 &#176;C.</li>
		<li>Apply eye lubricant, shave the pubic to the epigastric area, and cleanse the area with an iodine-based scrub and 70% alcohol.</li>
	</ol>
	</li>
	<li>Medications
	<ol>
		<li>Inject 2 mL of warm saline subcutaneously to compensate for the fluid lost during the surgery. Inject 200 IU of heparin subcutaneously.</li>
		<li>Administer antibiotic prophylaxis by injecting 10 mg/kg cefazolin dissolved in 0.3 mL of saline subcutaneously or intramuscularly.</li>
		<li>Administer pain control by injecting 20 mg/kg of diclofenac subcutaneously.</li>
	</ol>
	</li>
	<li>Perform the mid-line laparotomy and insert a retractor to widen the abdominal cavity. Mobilize the abdominal organs to the left side of the recipient using cotton swabs to make space for the procedure.</li>
	<li>Prevent dehydration by wrapping the abdominal organs with warm and wet gauze. Intermittingly spread warm saline with a 50 mL syringe during the surgery.</li>
	<li>Utilizing a surgical microscope with a 10x magnification, mobilize the duodenum and proximal jejunum by blunt dissection with cotton swabs to expose the Abd. A. and IVC. Prepare the Abd. A and IVC for anastomosis and systematically implant the donor heart, in accordance with Figure 3 or previously documented methods15. 
	NOTE: Do not separate the Abd. A. and IVC.
	<ol>
		<li>Assuming vascular anastomosis to be placed infrarenal, prepare a sufficient portion of the aorta and IVC for clamping.</li>
		<li>Perform blunt preparation using cotton swabs or sharp-serrated forceps to remove the fats and fascia around the vessels.</li>
		<li>Place 5-0 silk ligatures to the mesenteric branches and both the cranial and caudal sides of the major vessels. Elevate the abdominal vessels and coagulate or ligate the lumbar branches with 5-0 silk sutures. Remember to spare the testicular arteries and veins and do not clamp them.</li>
		<li>Use ligatures to lift the vessels and position the micro-clamps to the mesenteric branches, caudal, and cranial sides of the major vessels to stop the blood flow at the anastomosis site. Switch off the heating pad before placing the clamps, as excess heating can exacerbate limb ischemia. Ensure to switch on the heating pad after de-clamping the vessels to avoid hypothermia.</li>
		<li>Puncture the aorta using a 27 G needle and elongate the incision with micro scissors to a length equal to or slightly larger than the opening of the donor ascending aorta (Asc. A), which is approximately 5 mm.</li>
		<li>Make a longitudinal incision on the IVC in the same way as the aortotomy, but make it 3 mm closer to the caudal side compared to the aorta incision.</li>
		<li>Starting the anastomoses, placed the donor heart on the right side of the recipient&#39;s abdomen and attach the donor Asc. A to the recipient&#39;s Abd. A with one simple interrupted stitch (9-0 polypropylene) at the cranial corner of the longitudinal incision.</li>
		<li>Move the heart to the left side of the recipient abdomen and perform anastomosis of the donor&#39;s Asc. A with the recipient&#39;s Abd. A using a running 9-0 polypropylene suture.</li>
		<li>Fixate the donor pulmonary artery to the IVC with two interrupted sutures (9-0 polypropylene) at the caudal and cranial corners of the longitudinal incision.</li>
		<li>Perform the first half of the venous anastomosis from the intraluminal side of the vessel and complete the second half from the extraluminal side of the vessel. Before tightening the knots, flush the field with saline to prevent air embolism.</li>
	</ol>
	</li>
	<li>De-airing and de-clamping
	<ol>
		<li>Remove the mesenteric vein clamp first after completing the anastomosis to allow the right side of the heart to fill with venous blood.</li>
		<li>Remove the air in the coronary circuit and Asc. A. by applying retrograde coronary perfusion for several seconds.</li>
		<li>Place a piece of gauze on both sides of the vessels and remove the caudal clamp and the cranial clamp.</li>
		<li>Apply gentle compression with cotton swabs for 1-2 min. After ensuring adequate hemostasis, remove the swabs and wash the anastomoses with warm saline. 
		NOTE: The heart should begin beating within the first minute of reperfusion. If the recipient rat&#39;s body temperature is below 35 &#176;C, the heart rhythm will normalize after the temperature reaches 36 &#176;C.</li>
	</ol>
	</li>
	<li>Replace the abdominal organs in a meander-like manner and close the layers of the abdominal incision using continuous 5-0 polypropylene sutures.</li>
	<li>After the surgery, place the anesthetized animal on a clean area over a heating pad until the body temperature reaches 37&#176;C.&#160; 
	NOTE: Do not initiate the postoperative examinations till the body temperature reaches 37&#176;C. Maintain anesthesia at 2-2.5% isoflurane until the end of the experiments.</li>
	<li>Monitor the ECG of the transplanted donor heart for 3 h. Then, excise the heart under deep anesthesia for histological studies. 
	NOTE: Confirm anesthesia depth via lack of pedal reflex before excising the heart. The surgical procedure and the ECG monitoring take less than 6 h. Diclofenac, administered perioperatively (step 4.2.3.), enables pain management for the entire duration of this procedure. The analgesia regimen can be adjusted per the institutional animal use guidelines.</li>
</ol>

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J., 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=Normothermic+ex-situ+heart+perfusion+with+the+organ+care+system+for+cardiac+transplantation:+A+meta-analysis.">Normothermic ex-situ heart perfusion with the organ care system for cardiac transplantation: A meta-analysis.</a> Transplantation. 106 (9), 1745-1753 (2022).</li><li>Ardehali, A., 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=Ex-vivo+perfusion+of+donor+hearts+for+human+heart+transplantation+(PROCEED+II):+a+prospective,+open-label,+multicentre,+randomized+non-inferiority+trial.">Ex-vivo perfusion of donor hearts for human heart transplantation (PROCEED II): a prospective, open-label, multicentre, randomized non-inferiority trial.</a> Lancet. 385 (9987), 2577-2584 (2015).</li><li>Dang Van, S., 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=Ex+vivo+perfusion+of+the+donor+heart:+Preliminary+experience+in+high-risk+transplantations.">Ex vivo perfusion of the donor heart: Preliminary experience in high-risk transplantations.</a> Archives of Cardiovascular Diseases. 114 (11), 715-726 (2021).</li><li>Zhou, P., 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=Donor+heart+preservation+with+hypoxic-conditioned+medium-derived+from+bone+marrow+mesenchymal+stem+cells+improves+cardiac+function+in+a+heart+transplantation+model.">Donor heart preservation with hypoxic-conditioned medium-derived from bone marrow mesenchymal stem cells improves cardiac function in a heart transplantation model.</a> Stem Cell Research and Therapy. 12 (1), 5f6 (2021).</li><li>Messer, S., Large, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Resuscitating+heart+transplantation:+the+donation+after+circulatory+determined+death+donor.European.">Resuscitating heart transplantation: the donation after circulatory determined death donor.European.</a> Journal of Cardio-Thoracic Surgery. 49 (1), 1-4 (2016).</li><li>Trahanas, J. 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(188), e63945 (2022).</li><li>Fu, X., Segiser, A., Carrel, T. P., Tevaearai Stahel, H. 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The authors have no conflicts of interest.

Our focus in establishing this model was to replicate normothermic human heart transplantation. Non-ejecting models are the commonly preferred technique for preserving the donor heart in an ex situ environment16. While ejecting models offer many advantages in assessing cardiac function during ex situ perfusion17, they are not suitable for heterotopic transplantation models. In heterotopic transplantation, the implanted donor heart needs to overcome systoli...

An erratum was issued for:&#160;<a href="https://www.jove.com/t/64954">Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation</a>. The Protocol section was updated.
Section 4 of the Protocol was updated from:
4. Implantation
<ol>
	<li>Preparation of recipient
	<ol>
		<li>Begin the recipient preparation 30 min before the cessation of ex situ perfusion.</li>
		<li>Anesthetize the recipient animal using the same method as mentioned in step 2.2.</li>
		<li>Place the rat in a supine position on the heating pad and insert the temperature probe into the rectum to maintain the body temperature at 37 &#176;C.</li>
		<li>Apply eye lubricant, shave the pubic to the epigastric area, and cleanse the area with an iodine-based scrub and 70% alcohol.</li>
	</ol>
	</li>
	<li>Medications
	<ol>
		<li>Inject 2 mL of warm saline subcutaneously to compensate for the fluid lost during the surgery. Inject 200 IU of heparin subcutaneously.</li>
		<li>Administer antibiotic prophylaxis by injecting 10 mg/kg cefazolin dissolved in 0.3 mL of saline subcutaneously or intramuscularly.</li>
		<li>Administer pain control by injecting 20 mg/kg of diclofenac subcutaneously.</li>
	</ol>
	</li>
	<li>Perform the mid-line laparotomy and insert a retractor to widen the abdominal cavity. Mobilize the abdominal organs to the left side of the recipient using cotton swabs to make space for the procedure.</li>
	<li>Prevent dehydration by wrapping the abdominal organs with warm and wet gauze. Intermittingly spread warm saline with a 50 mL syringe during the surgery.</li>
	<li>Utilizing a surgical microscope with a 10x magnification, mobilize the duodenum and proximal jejunum by blunt dissection with cotton swabs to expose the Abd. A. and IVC. Prepare the Abd. A and IVC for anastomosis and systematically implant the donor heart, in accordance with Figure 3 or previously documented methods15. 
	NOTE: Do not separate the Abd. A. and IVC.
	<ol>
		<li>Assuming vascular anastomosis to be placed infrarenal, prepare a sufficient portion of the aorta and IVC for clamping.</li>
		<li>Perform blunt preparation using cotton swabs or sharp-serrated forceps to remove the fats and fascia around the vessels.</li>
		<li>Place 5-0 silk ligatures to the mesenteric branches and both the cranial and caudal sides of the major vessels. Elevate the abdominal vessels and coagulate or ligate the lumbar branches with 5-0 silk sutures. Remember to spare the testicular arteries and veins and do not clamp them.</li>
		<li>Use ligatures to lift the vessels and position the micro-clamps to the mesenteric branches, caudal, and cranial sides of the major vessels to stop the blood flow at the anastomosis site. Be sure to switch off the heating pad before placing the clamps, as excess heating can exacerbate limb ischemia.</li>
		<li>Puncture the aorta using a 27 G needle and elongate the incision with micro scissors to a length equal to or slightly larger than the opening of the donor ascending aorta (Asc. A), which is approximately 5 mm.</li>
		<li>Make a longitudinal incision on the IVC in the same way as the aortotomy, but make it 3 mm closer to the caudal side compared to the aorta incision.</li>
		<li>Starting the anastomoses, placed the donor heart on the right side of the recipient&#39;s abdomen and attach the donor Asc. A to the recipient&#39;s Abd. A with one simple interrupted stitch (9-0 polypropylene) at the cranial corner of the longitudinal incision.</li>
		<li>Move the heart to the left side of the recipient abdomen and perform anastomosis of the donor&#39;s Asc. A with the recipient&#39;s Abd. A using a running 9-0 polypropylene suture.</li>
		<li>Fixate the donor pulmonary artery to the IVC with two interrupted sutures (9-0 polypropylene) at the caudal and cranial corners of the longitudinal incision.</li>
		<li>Perform the first half of the venous anastomosis from the intraluminal side of the vessel and complete the second half from the extraluminal side of the vessel. Before tightening the knots, flush the field with saline to prevent air embolism.</li>
	</ol>
	</li>
	<li>De-airing and de-clamping
	<ol>
		<li>Remove the mesenteric vein clamp first after completing the anastomosis to allow the right side of the heart to fill with venous blood.</li>
		<li>Remove the air in the coronary circuit and Asc. A. by applying retrograde coronary perfusion for several seconds.</li>
		<li>Place a piece of gauze on both sides of the vessels and remove the caudal clamp and the cranial clamp.</li>
		<li>Apply gentle compression with cotton swabs for 1-2 min. After ensuring adequate hemostasis, remove the swabs and wash the anastomoses with warm saline. 
		NOTE: The heart should begin beating within the first minute of reperfusion. If the recipient rat&#39;s body temperature is below 35 &#176;C, the heart rhythm will normalize after the temperature reaches 36 &#176;C.</li>
	</ol>
	</li>
	<li>Replace the abdominal organs in a meander-like manner and close the layers of the abdominal incision using continuous 5-0 polypropylene sutures.</li>
</ol>
to:
4. Implantation
<ol>
	<li>Preparation of recipient
	<ol>
		<li>Begin the recipient preparation 30 min before the cessation of ex situ perfusion.</li>
		<li>Anesthetize the recipient animal using the same method as mentioned in step 2.2.</li>
		<li>Place the rat in a supine position on the heating pad and insert the temperature probe into the rectum to maintain the body temperature at 37 &#176;C.</li>
		<li>Apply eye lubricant, shave the pubic to the epigastric area, and cleanse the area with an iodine-based scrub and 70% alcohol.</li>
	</ol>
	</li>
	<li>Medications
	<ol>
		<li>Inject 2 mL of warm saline subcutaneously to compensate for the fluid lost during the surgery. Inject 200 IU of heparin subcutaneously.</li>
		<li>Administer antibiotic prophylaxis by injecting 10 mg/kg cefazolin dissolved in 0.3 mL of saline subcutaneously or intramuscularly.</li>
		<li>Administer pain control by injecting 20 mg/kg of diclofenac subcutaneously.</li>
	</ol>
	</li>
	<li>Perform the mid-line laparotomy and insert a retractor to widen the abdominal cavity. Mobilize the abdominal organs to the left side of the recipient using cotton swabs to make space for the procedure.</li>
	<li>Prevent dehydration by wrapping the abdominal organs with warm and wet gauze. Intermittingly spread warm saline with a 50 mL syringe during the surgery.</li>
	<li>Utilizing a surgical microscope with a 10x magnification, mobilize the duodenum and proximal jejunum by blunt dissection with cotton swabs to expose the Abd. A. and IVC. Prepare the Abd. A and IVC for anastomosis and systematically implant the donor heart, in accordance with Figure 3 or previously documented methods15. 
	NOTE: Do not separate the Abd. A. and IVC.
	<ol>
		<li>Assuming vascular anastomosis to be placed infrarenal, prepare a sufficient portion of the aorta and IVC for clamping.</li>
		<li>Perform blunt preparation using cotton swabs or sharp-serrated forceps to remove the fats and fascia around the vessels.</li>
		<li>Place 5-0 silk ligatures to the mesenteric branches and both the cranial and caudal sides of the major vessels. Elevate the abdominal vessels and coagulate or ligate the lumbar branches with 5-0 silk sutures. Remember to spare the testicular arteries and veins and do not clamp them.</li>
		<li>Use ligatures to lift the vessels and position the micro-clamps to the mesenteric branches, caudal, and cranial sides of the major vessels to stop the blood flow at the anastomosis site. Switch off the heating pad before placing the clamps, as excess heating can exacerbate limb ischemia. Ensure to switch on the heating pad after de-clamping the vessels to avoid hypothermia.</li>
		<li>Puncture the aorta using a 27 G needle and elongate the incision with micro scissors to a length equal to or slightly larger than the opening of the donor ascending aorta (Asc. A), which is approximately 5 mm.</li>
		<li>Make a longitudinal incision on the IVC in the same way as the aortotomy, but make it 3 mm closer to the caudal side compared to the aorta incision.</li>
		<li>Starting the anastomoses, placed the donor heart on the right side of the recipient&#39;s abdomen and attach the donor Asc. A to the recipient&#39;s Abd. A with one simple interrupted stitch (9-0 polypropylene) at the cranial corner of the longitudinal incision.</li>
		<li>Move the heart to the left side of the recipient abdomen and perform anastomosis of the donor&#39;s Asc. A with the recipient&#39;s Abd. A using a running 9-0 polypropylene suture.</li>
		<li>Fixate the donor pulmonary artery to the IVC with two interrupted sutures (9-0 polypropylene) at the caudal and cranial corners of the longitudinal incision.</li>
		<li>Perform the first half of the venous anastomosis from the intraluminal side of the vessel and complete the second half from the extraluminal side of the vessel. Before tightening the knots, flush the field with saline to prevent air embolism.</li>
	</ol>
	</li>
	<li>De-airing and de-clamping
	<ol>
		<li>Remove the mesenteric vein clamp first after completing the anastomosis to allow the right side of the heart to fill with venous blood.</li>
		<li>Remove the air in the coronary circuit and Asc. A. by applying retrograde coronary perfusion for several seconds.</li>
		<li>Place a piece of gauze on both sides of the vessels and remove the caudal clamp and the cranial clamp.</li>
		<li>Apply gentle compression with cotton swabs for 1-2 min. After ensuring adequate hemostasis, remove the swabs and wash the anastomoses with warm saline. 
		NOTE: The heart should begin beating within the first minute of reperfusion. If the recipient rat&#39;s body temperature is below 35 &#176;C, the heart rhythm will normalize after the temperature reaches 36 &#176;C.</li>
	</ol>
	</li>
	<li>Replace the abdominal organs in a meander-like manner and close the layers of the abdominal incision using continuous 5-0 polypropylene sutures.</li>
	<li>After the surgery, place the anesthetized animal on a clean area over a heating pad until the body temperature reaches 37&#176;C.&#160; 
	NOTE: Do not initiate the postoperative examinations till the body temperature reaches 37&#176;C. Maintain anesthesia at 2-2.5% isoflurane until the end of the experiments.</li>
	<li>Monitor ECG of the transplanted donor heart for 3 h. Then, excise the heart under deep anesthesia for histological studies. 
	NOTE: Confirm anesthesia depth via lack of pedal reflex before excising the heart. The surgical procedure and the ECG monitoring take less than 6 h. Diclofenac, administered perioperatively (step 4.2.3.), enables pain management for the entire duration of this procedure. The analgesia regimen can be adjusted per the institutional animal use guidelines.</li>
</ol>

An erratum was issued for:&#160;<a href="https://www.jove.com/t/64954">Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation</a>. The Protocol section was updated.

Erratum: Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation

Heart transplantation remains the sole viable therapy for refractory heart failure1,2,3,4. Despite a steady rise in the number of patients in need of heart transplantation, a proportional increase in the availability of donor organs has not been observed5. To address this issue, novel approaches for preserving donor hearts have been developed with the goal of improving the challenges and increasing the availability of donors6,7,8,9.
Normothermic ex situ heart perfusion (NESHP) using organ care system (OCS) machines has emerged as a clinical intervention1,3. This technique has been deemed a suitable alternative to the conventional static cold storage (SCS) method2,9. NESHP effectively reduces the duration of cold ischemia, diminishes metabolic demand, and facilitates optimal nutritional supply and oxygenation during the transportation of donor organs10,11. Despite the clear potential of this method to improve donor organ preservation, its clinical application and further investigation have been constrained by high costs. Therefore, preclinical animal models of NESHP are crucial for identifying key technical challenges associated with this technique12,13. Pigs and rats are the preferred animal models for preclinical studies due to their ischemic tolerance9. Although the porcine model is ideal for basic and translational research, it is limited by its high cost and the intensive labor required for care and maintenance. In contrast, rat models are less expensive and easier to handle14.
In this study, we introduce a simplified rat model of NESHP, followed by heterotopic heart transplantation, to evaluate the impact of the preservation technique on graft condition post-implantation. This model is straightforward, cost-effective, and can be executed by a single experimenter. Figure 1 shows the schematics of the procedure.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>AES active evacuation system</td>
			<td>Smiths medical</td>
			<td>PC-6769-51A</td>
			<td>Utilize CO2 and excess isoflurane</td>
		</tr>
		<tr>
			<td>Anesthesia machine</td>
			<td>Smiths medical</td>
			<td>PC-8801-01A</td>
			<td>Mixes isoflurane and oxyegn and delivers to animal</td>
		</tr>
		<tr>
			<td>B20 patient monitor</td>
			<td>GE medical systems</td>
			<td>B20</td>
			<td>to observe mean aortic pressure and temperature</td>
		</tr>
		<tr>
			<td>Homeothermic Monitoring System</td>
			<td>Harvard apparatus</td>
			<td>55-7020</td>
			<td>To monitor and maintain animal&#39;s temperature</td>
		</tr>
		<tr>
			<td>Micro-1 Rat oxygenator</td>
			<td>Dongguan Kewei medical instruments</td>
			<td>Micro-MO</td>
			<td>For gas exchange in the langendorff circuit</td>
		</tr>
		<tr>
			<td>Micropuncture introducer Set</td>
			<td>COOK medical</td>
			<td>G48007</td>
			<td>for delivering cardioplegic solution to the arch through the abdominal aorta</td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Amscope</td>
			<td>MU1403</td>
			<td>For zooming surgical field (Recipient)</td>
		</tr>
		<tr>
			<td>Surgical loupe</td>
			<td>SurgiTel</td>
			<td>L2S09</td>
			<td>For zooming surgical field (Donor)</td>
		</tr>
		<tr>
			<td>Syringe pump</td>
			<td>AMP all</td>
			<td>SP-8800</td>
			<td>To deliver cardioplegic solution</td>
		</tr>
		<tr>
			<td>Transonic flow sensor</td>
			<td>Transonic</td>
			<td>ME3PXL-M5</td>
			<td>Perfusion circuit flow sensor</td>
		</tr>
		<tr>
			<td>Transonic tubing flow module</td>
			<td>Transonic</td>
			<td>TS410</td>
			<td>flow acquiring system</td>
		</tr>
		<tr>
			<td>Watson - Marlow pumps</td>
			<td>Harvard apparatus</td>
			<td>010.6131.DAO</td>
			<td>Peristaltic pump used for recirculate perfusate</td>
		</tr>
		<tr>
			<td>WBC-1510A</td>
			<td>JEIO TECH</td>
			<td>E03056D</td>
			<td>Heating bath</td>
		</tr>
		<tr>
			<td>Sprague-Dawley rats</td>
			<td>Samtako Bio Korea Co., Ltd., Osan City Korea</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Medications</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BioHAnce Gel Eye Drops</td>
			<td>SENTRIX Animal care</td>
			<td></td>
			<td>wet ointments for eye</td>
		</tr>
		<tr>
			<td>Cefazolin</td>
			<td>JW pharmaceutical</td>
			<td></td>
			<td>For prophilaxis</td>
		</tr>
		<tr>
			<td>Custodiol</td>
			<td>DR, FRANZ KOHLER CHEMIE GMBH</td>
			<td></td>
			<td>For heart harvesting</td>
		</tr>
		<tr>
			<td>Diclofenac</td>
			<td>Myungmoon Pharm. Co. Ltd</td>
			<td></td>
			<td>For pain control</td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>JW pharmaceutical</td>
			<td></td>
			<td>Anticoagulant</td>
		</tr>
		<tr>
			<td>Insulin</td>
			<td>JW pharmaceutical</td>
			<td></td>
			<td>hormon therapy</td>
		</tr>
		<tr>
			<td>Saline</td>
			<td>JW pharmaceutical</td>
			<td></td>
			<td>For hydration therapy</td>
		</tr>
	</tbody>
</table>

rat model of normothermic ex-situ perfused heterotopic heart transplantation

Heart transplantation is the most effective therapy for end-stage heart failure. Despite the improvements in therapeutic approaches and interventions, the number of heart failure patients waiting for transplantation is still increasing. The normothermic ex situ preservation technique has been established as a comparable method to the conventional static cold storage technique. The main advantage of this technique is that donor hearts can be preserved for up to 12 h in a physiologic condition. Moreover, this technique allows resuscitation of the donor hearts after circulatory death and applies required pharmacologic interventions to improve donor function after implantation. Numerous animal models have been established to improve normothermic ex situ preservation techniques and eliminate preservation-related complications. Although large animal models are easy to handle compared to small animal models, it is costly and challenging. We present a rat model of normothermic ex situ donor heart preservation followed by heterotopic abdominal transplantation. This model is relatively cheap and can be accomplished by a single experimenter.

Figure 1 illustrates the experimental design used in a small animal model. Figure 2 displays the modified Langendorff perfusion apparatus, which includes a small animal oxygenator. The order of anastomosis for heterotopic abdominal implantation is presented in Figure 3.
Figure 4 shows the parameters used to assess the viability of the heart during ex situ perfusion, ...

Watch this Scientific Journal Video about Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation at JoVE.com

Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation

Here, we present an assessment protocol of a heterotopically implanted heart after normothermic ex situ preservation in the rat model.

Heart transplantation is the most effective therapy for end-stage heart failure. Despite the improvements in therapeutic approaches and interventions, the ...

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951 Views.  Chonnam National University Graduate School. Here, we present an assessment protocol of a heterotopically implanted heart after normothermic ex situ preservation in the rat model.

Video: Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation

Murine Heterotopic Heart Transplant Technique

It is now over forty years since this technique was first reported by Corry, Wynn and Russell. Although it took some years for other labs to become proficient in and utilize this technique, it is now widely used by many laboratories around the world. A significant refinement to the original technique was developed and reported in 2001 by Niimi. Described here are the techniques that have evolved over more than a decade in the hands of three surgeons (Plenter, Grazia, Pietra) in our center. These techniques are now being passed on to a younger generation of surgeons and researchers.
Based largely on the Niimi experience, the procedures used have evolved in the fine details - details which we will endeavor to relate here in such a way that others may be able to use this very useful model. Like Niimi, we have found that a video aid to learning is a priceless resource for the beginner.

The manuscript describes the steps required to perform the heterotopic heart transplant in the mouse.

It is now over forty years since this technique was first reported by Corry, Wynn and Russell. Although it took some years for other labs to become ...

Transplantation of Pulmonary Valve Using a Mouse Model of Heterotopic Heart Transplantation

Tissue engineered heart valves, especially decellularized valves, are starting to gain momentum in clinical use of reconstructive surgery with mixed results. However, the cellular and molecular mechanisms of the neotissue development, valve thickening, and stenosis development are not researched extensively. To answer the above questions, we developed a murine heterotopic heart valve transplantation model. A heart valve was harvested from a valve donor mouse and transplanted to a heart donor mouse. The heart with a new valve was transplanted heterotopically to a recipient mouse. The transplanted heart showed its own heartbeat, independent of the recipient&#8217;s heartbeat. The blood flow was quantified using a high frequency ultrasound system with a pulsed wave Doppler. The flow through the implanted pulmonary valve showed forward flow with minimal regurgitation and the peak flow was close to 100 mm/sec. This murine model of heart valve transplantation is highly versatile, so it can be modified and adapted to provide different hemodynamic environments and/or can be used with various transgenic mice to study neotissue development in a tissue engineered heart valve.

In order to understand the cellular and molecular mechanisms underlying neotissue formation and stenosis development in tissue engineered heart valves, a murine model of heterotopic heart valve transplantation was developed. A pulmonary heart valve was transplanted to recipient using the heterotopic heart transplantation technique.

Tissue engineered heart valves, especially decellularized valves, are starting to gain momentum in clinical use of reconstructive surgery with mixed ...

Rat Heterotopic Abdominal Heart/Single-lung Transplantation in a Volume-loaded Configuration

Herein, we describe a novel technique for heterotopic abdominal heart-lung transplantation (HAHLT) in rats. The configuration of the transplant graft involves anastomosis of donor inferior vena cava (IVC) to recipient IVC, and donor ascending aorta (Ao) to recipient abdominal Ao. The right upper and middle lung lobes are preserved and function as conduits for blood flow from right heart to left heart.
There are several advantages to using this technique, and it lends itself to a broad range of applications. Because the graft is transplanted in a configuration that allows for dyamic volume-loading, cardiac function may be directly assessed in vivo. The use of pressure-volume conductance catheters permits characterization of load-dependent and load-independent hemodynamic parameters. The graft may be converted to a loaded configuration by applying a clamp to the recipient&#8217;s infra-hepatic IVC. We describe modified surgical techniques for both donor and recipient operations, and an ideal myocardial protection strategy. Depending on the experimental aim, this model may be adapted for use in both acute and chronic studies of graft function, immunologic status, and variable ventricular loading conditions. The conducting airways to the transplanted lung are preserved, and allow for acute lung re-ventilation. This facilitates analysis of the effects of the mixed venous and arterial blood providing coronary perfusion to the graft.
A limitation of this model is its technical complexity. There is a significant learning curve for new operators, who should ideally be mentored in the technique. A surgical training background is advantageous for those wishing to apply this model. Despite its complexity, we aim to present the model in a clear and easily applicable format. Because of the physiologic similarity of this model to orthotopic transplantation, and its broad range of study applications, the effort invested in learning the technique is likely to be worthwhile.

We describe a novel technique for heterotopic abdominal heart-lung transplantation (HAHLT) in rats. The transplant configuration results in a partially loaded graft circulation, allowing direct functional assessment. This model may be employed for acute or chronic studies of function and immunologic status of the transplanted graft.

Herein, we describe a novel technique for heterotopic abdominal heart-lung transplantation (HAHLT) in rats. The configuration of the transplant graft ...

Ex Situ Normothermic Machine Perfusion of Donor Livers

In contrast to conventional static cold preservation (0-4 &#176;C), ex situ machine perfusion may provide better preservation of donor livers. Continuous perfusion of organs provides the opportunity to improve organ quality and allows ex situ viability assessment of donor livers prior to transplantation. This video article provides a step by step protocol for ex situ normothermic machine perfusion (37 &#176;C) of human donor livers using a device that provides a pressure and temperature controlled pulsatile perfusion of the hepatic artery and continuous perfusion of the portal vein. The perfusion fluid is oxygenated by two hollow fiber membrane oxygenators and the temperature can be regulated between 10 &#176;C and 37 &#176;C. During perfusion, the metabolic activity of the liver as well as the degree of injury can be assessed by biochemical analysis of samples taken from the perfusion fluid. Machine perfusion is a very promising tool to increase the number of livers that are suitable for transplantation.

Ex Situ Normothermic Machine Perfusion of Donor Livers

Here we present a protocol describing oxygenated ex situ machine perfusion of donor liver grafts. This article contains a step by step protocol to procure and prepare the liver graft for machine perfusion, prepare the perfusion fluid, prime the perfusion machine and perform oxygenated normothermic machine perfusion of the liver graft.

In contrast to conventional static cold preservation (0-4 &#176;C), ex situ machine perfusion may provide better preservation of donor livers. Continuous ...

Heterotopic Cervical Heart Transplantation in Mice

The heterotopic cervical heart transplantation in mice is a valuable tool in transplant and cardiovascular research. The cuff technique greatly simplifies this model by avoiding challenging suture anastomoses of small vessels thereby reducing warm ischemia time. In comparison to abdominal graft implantation the cervical model is less invasive and the implanted graft is easily accessible for further follow-up examinations. Anastomoses are performed by pulling the ascending aorta of the graft over the cuff with the recipient&#8217;s common carotid artery and by pulling the main pulmonary artery over the cuff with the external jugular vein. Selection of appropriate cuff size and complete mobilization of the vessels are important for successful revascularization. Ischemia-reperfusion (I/R) injury can be minimized by perfusing the graft with a cardioplegic solution and by hypothermia. In this article, we provide technical details for a simplified and improved cuff technique, which should allow surgeons with basic microsurgical skills to perform the procedure with a high success rate.

The cuff technique shortens and facilitates the model of heterotopic cervical heart transplantation in mice by avoiding technically challenging suture anastomoses of small vessels. The technical details shown in this video paper should allow researches to establish this model in their laboratories.

The heterotopic cervical heart transplantation in mice is a valuable tool in transplant and cardiovascular research. The cuff technique greatly simplifies ...

Normothermic Ex Situ Heart Perfusion in Working Mode: Assessment of Cardiac Function and Metabolism

The current standard method for organ preservation (cold storage, CS), exposes the heart to a period of cold ischemia that limits the safe preservation time and increases the risk of adverse post-transplantation outcomes. Moreover, the static nature of CS does not allow for organ evaluation or intervention during the preservation interval. Normothermic ex situ heart perfusion (ESHP) is a novel method for preservation of the donated heart that minimizes cold ischemia by providing oxygenated, nutrient-rich perfusate to the heart. ESHP has been shown to be non-inferior to CS in the preservation of standard-criteria donor hearts and has also facilitated the clinical transplantation of the hearts donated after the circulatory determination of death. Currently, the only available clinical ESHP device perfuses the heart in an unloaded, non-working state, limiting assessments of myocardial performance. Conversely, ESHP in working mode provides the opportunity for comprehensive evaluation of cardiac performance by assessment of functional and metabolic parameters under physiologic conditions. Moreover, earlier experimental studies have suggested that ESHP in working mode may result in improved functional preservation. Here, we describe the protocol for ex situ perfusion of the heart in a large mammal (porcine) model, which is reproducible for different animal models and heart sizes. The software program in this ESHP apparatus allows for real-time and automated control of the pump speed to maintain desired aortic and left atrial pressure and evaluates a variety of functional and electrophysiological parameters with minimal need for supervision/manipulation.

Normothermic ex situ heart perfusion (ESHP), preserves the heart in a beating, semi-physiologic state. When performed in a working mode, ESHP provides the opportunity to perform sophisticated assessments of donor heart function and organ viability. Here, we describe our method for myocardial performance evaluation during ESHP.

The current standard method for organ preservation (cold storage, CS), exposes the heart to a period of cold ischemia that limits the safe preservation ...

A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of transplant with the potential to deliver durable heart valves capable of somatic growth with no requirement for anticoagulation. However, the immunobiology of heart valve transplants remains unexplored, highlighting the need for animal models to study this new type of transplant. Previous rat models for heterotopic aortic valve transplantation into the abdominal aorta have been described, though they are technically challenging and costly. For addressing this challenge, a renal subcapsular transplant model was developed in rodents as a practical and more straightforward method for studying heart valve transplant immunobiology. In this model, a single aortic valve leaflet is harvested and inserted into the renal subcapsular space. The kidney is easily accessible, and the transplanted tissue is securely contained in a subcapsular space that is well vascularized and can accommodate a variety of tissue sizes. Furthermore, because a single rat can provide three donor aortic leaflets and a single kidney can provide multiple sites for transplanted tissue, fewer rats are required for a given study. Here, the transplantation technique is described, providing a significant step forward in studying the transplant immunology of heart valve transplantation.

This protocol describes a simple and efficient method for the transplantation of aortic valve leaflets under the renal capsule to allow for the study of alloreactivity of heart valves.

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of ...

Normothermic Negative Pressure Ventilation Ex Situ Lung Perfusion: Evaluation of Lung Function and Metabolism

Lung transplantation (LTx) remains the standard of care for end-stage lung disease. A shortage of suitable donor organs and concerns over donor organ quality exacerbated by excessive geographic transportation distance and stringent donor organ acceptance criteria pose limitations to current LTx efforts. Ex situ lung perfusion (ESLP) is an innovative technology that has shown promise in attenuating these limitations. The physiologic ventilation and perfusion of the lungs outside of the inflammatory milieu of the donor body affords ESLP several advantages over traditional cold static preservation (CSP). There is evidence that negative pressure ventilation (NPV) ESLP is superior to positive pressure ventilation (PPV) ESLP, with PPV inducing more significant ventilator-induced lung injury, pro-inflammatory cytokine production, pulmonary edema, and bullae formation. The NPV advantage is perhaps due to the homogenous distribution of intrathoracic pressure across the entire lung surface. The clinical safety and feasibility of a custom NPV-ESLP device have been demonstrated in a recent clinical trial involving extender criteria donor (ECD) human lungs. Herein, the use of this custom device is described in a juvenile porcine model of normothermic NPV-ESLP over a 12 h duration, paying particular attention to management techniques. Pre-surgical preparation, including ESLP software initialization, priming, and de-airing of the ESLP circuit, and the addition of anti-thrombotic, anti-microbial, and anti-inflammatory agents, is specified. The intraoperative techniques of central line insertion, lung biopsy, exsanguination, blood collection, cardiectomy, and pneumonectomy are described. Furthermore, particular focus is paid to anesthetic considerations, with anesthesia induction, maintenance, and dynamic modifications outlined. The protocol also specifies the custom device's initialization, maintenance, and termination of perfusion and ventilation. Dynamic organ management techniques, including alterations in ventilation and metabolic parameters to optimize organ function, are thoroughly described. Finally, the physiological and metabolic assessment of lung function is characterized and depicted in the representative results.

Normothermic Negative Pressure Ventilation Ex Situ Lung Perfusion: Evaluation of Lung Function and Metabolism

This paper describes a porcine model of negative pressure ventilation ex situ lung perfusion, including procurement, attachment, and management on the custom-made platform. Focus is made on anesthetic and surgical techniques, as well as troubleshooting.

Lung transplantation (LTx) remains the standard of care for end-stage lung disease. A shortage of suitable donor organs and concerns over donor organ ...

A Modified Cuff Technique for Mouse Cervical Heterotopic Heart Transplantation Model

Cardiac allograft rejection limits the long-term survival of patients after heart transplantation. A mouse heart transplantation model is ideal for investigating the mechanism of cardiac allograft rejection in preclinical studies because of their high homology with human genes. This understanding would help develop unique approaches to improving patients&#39; long-term survival treated with cardiac allografts. In a mouse model, abdominal donor heart implantation is commonly performed with an end-to-end&#160;anastomosis to the recipient&#39;s aorta and inferior vena cava using stitches. In this model, the donor&#39;s heart is implanted by end-to-end anastomosis to the recipient&#39;s carotid artery and jugular vein by the modified-Cuff technique. The transplantation surgery is performed without stitching and thus may increase the survival of the recipient since there is no interference with the blood supply and venous reflux of the lower body. This mouse model would help investigate the mechanisms underlying the immunological and pathological (acute/chronic) rejection of cardiac allografts.

In the present protocol, a mouse heart transplantation model is used for investigating the mechanism of cardiac allograft rejection. In this heterotopic heart transplantation model, operation efficiency is improved, and the survival of cardiac grafts is ensured by a cervical end-to-end anastomosis of heart implantation using a modified Cuff technique.

Cardiac allograft rejection limits the long-term survival of patients after heart transplantation. A mouse heart transplantation model is ideal for ...

Modified Heterotopic Abdominal Heart Transplantation and a Novel Aortic Regurgitation Model in Rats

Over the past 50 years, many researchers have reported heterotopic abdominal heart transplantation in mice and rats, with some variations in the surgical technique. Modifying the transplantation procedure to strengthen the myocardial protection could prolong the ischemia time while preserving the donor&#39;s cardiac function. This technique&#39;s key points are as follows: transecting the donor&#39;s abdominal aorta before harvesting to unload the donor&#39;s heart; perfusing the donor&#39;s coronary arteries with a cold cardioplegic solution; and topical cooling of the donor&#39;s heart during the anastomosis procedure. Consequently, since this procedure prolongs the acceptable ischemia time, beginners can easily perform it and achieve a high success rate.
Moreover, a new aortic regurgitation (AR) model was established in this work using a technique different from the existing one,&#160;which is created by inserting a catheter from the right carotid artery and puncturing the native aortic valve under continuous echocardiographic guidance. A heterotopic abdominal heart transplantation was performed using the novel AR model. In the protocol, after the donor&#39;s heart is harvested, a stiff guidewire is inserted into the donor&#39;s brachiocephalic artery and advanced toward the aortic root. The aortic valve is punctured by pushing the guidewire further even after the resistance is felt, thus inducing AR. It is easier to damage the aortic valve using this method than with the procedure described in the conventional AR model. Additionally, this novel AR model does not contribute to the recipient&#39;s circulation; therefore, this method is expected to produce a more severe AR model than the conventional procedure.

This study demonstrates a reproducible heterotopic abdominal heart transplantation technique in rats that beginners can learn and perform. Additionally, a novel aortic regurgitation model in rats is generated by performing heterotopic abdominal heart transplantation and damaging the donor's aortic valve using a guidewire after harvesting.

Over the past 50 years, many researchers have reported heterotopic abdominal heart transplantation in mice and rats, with some variations in the surgical ...