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><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&#039;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>&lt;strong&gt;Medications&lt;/strong&gt;</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, ...

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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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961 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

A Modified Method for Heterotopic Mouse Heart Transplantion

Mice are often used as heart transplant donors and recipients in studies of transplant immunology due to the wide range of transgenic mice and reagents available. A difficulty is presented due to the small size of the animal and the considerable technical challenges of the microsurgery involved in heart transplantation. In particular, a high rate of technical failure early after transplantation may result from recipient death and post-operative complications such as hind limb paralysis or a non-beating heart. Here, the complete technique for heterotopic mouse heart transplantation is demonstrated, involving harvesting the donor heart and its subsequent implantation into a recipient mouse. The donor heart is harvested immediately following in situ perfusion with cold heparinized saline and transection of the ascending aorta and pulmonary artery. The recipient operation involves preparation of the abdominal aorta and inferior vena cava (IVC), followed by end-to-side anastomosis of the donor aorta with the recipient aorta using a single running 10-0 microsuture and a similar anastomosis of the donor pulmonary artery with the recipient IVC. Following the operation the animal is injected with 0.6 ml normal saline subcutaneously and allowed to recover on a 37&#160;&#176;C heating pad. The results from 227 mouse heart transplants are summarized with a success rate at 48 hr of 86.8%. Of the 13.2% failures within 48 hr, 5 (2.2%) experienced hind limb paralysis, 10 (4.4%) had a non-beating heart due to graft ischemic injury and/or thrombosis, while 15 (6.6%) died within 48 hr.

A technique is demonstrated for the microsurgical procedure for heterotopic transplantation of hearts in mice, including simplified methods for donor harvesting and recipient vessel anastomosis.

Mice are often used as heart transplant donors and recipients in studies of transplant immunology due to the wide range of transgenic mice and reagents ...

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

This protocol presents an optimized erythrocytes-free NEVLP system using mouse livers. Ex vivo preservation of mouse livers was achieved by employing modified cannulas and techniques adapted from conventional commercial ex vivo perfusion equipment. The system was utilized to evaluate the preservation outcomes following 12 h of perfusion. C57BL/6J mice served as liver donors, and the livers were explanted by cannulating the portal vein (PV) and bile duct (BD), and subsequently flushing the organ with warm (37 &#176;C) heparinized saline. Then, the explanted livers were transferred to the perfusion chamber and subjected to normothermic oxygenated machine perfusion (NEVLP). Inlet and outlet perfusate samples were collected at 3 h intervals for perfusate analysis. Upon completion of the perfusion, liver samples were obtained for histological analysis, with morphological integrity assessed using modified Suzuki-Score through Hematoxylin-Eosin (HE) staining. The optimization experiments yielded the following findings: (1) mice weighing over 30 g were deemed more suitable for the experiment due to the larger size of their bile duct (BD). (2) a 2 Fr (outer diameter = 0.66 mm) polyurethane cannula was better suited for cannulating the portal vein (PV) when compared to a polypropylene cannula. This was attributed to the polyurethane material&#39;s enhanced grip, resulting in reduced catheter slippage during the transfer from the body to the organ chamber. (3) for cannulation of the bile duct (BD), a 1 Fr (outer diameter = 0.33 mm) polyurethane cannula was found to be more effective compared to the polypropylene UT - 03 (outer diameter = 0.30 mm) cannula. With this optimized protocol, mouse livers were successfully preserved for a duration of 12 h without significant impact on the histological structure. Hematoxylin-Eosin (HE) staining revealed a well-preserved morphological architecture of the liver, characterized by predominantly viable hepatocytes with clearly visible nuclei and mild dilation of hepatic sinusoids.

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

A normothermic ex vivo liver perfusion (NEVLP) system was created for mouse livers. This system requires experience in microsurgery but allows for reproducible perfusion results. The ability to utilize mouse livers facilitates the investigation of molecular pathways to identify novel perfusate additives and enables the execution of experiments focused on organ repair.

This protocol presents an optimized erythrocytes-free NEVLP system using mouse livers. Ex vivo preservation of mouse livers was achieved by employing ...

Engineering

Normothermic Ex Vivo Kidney Perfusion for the Preservation of Kidney Grafts prior to Transplantation

Kidney transplantation has become a well-established treatment option for patients with end-stage renal failure. The persisting organ shortage remains a serious problem. Therefore, the acceptance criteria for organ donors have been extended leading to the usage of marginal kidney grafts. These marginal organs tolerate cold storage poorly resulting in increased preservation injury and higher rates of delayed graft function. To overcome the limitations of cold storage, extensive research is focused on alternative normothermic preservation methods.
Ex vivo normothermic organ perfusion is an innovative preservation technique. The first experimental and clinical trials for ex vivo lung, liver, and kidney perfusions demonstrated favorable outcomes.
In addition to the reduction of cold ischemic injury, the method of normothermic kidney storage offers the opportunity for organ assessment and repair. This manuscript provides information about kidney retrieval, organ preservation techniques, and isolated ex vivo normothermic kidney perfusion (NEVKP) in a porcine model. Surgical techniques, set up for the perfusion solution and the circuit, potential assessment options, and representative results are demonstrated.

Normothermic Ex Vivo Kidney Perfusion for the Preservation of Kidney Grafts prior to Transplantation

The severe organ shortage has resulted in increased use of marginal kidney grafts for transplantation. This has triggered interest in alternative storage methods, since marginal grafts especially tolerate cold storage poorly. The technique of normothermic ex vivo kidney perfusion (NEVKP) represents a novel preservation method for kidney grafts prior to &#160;transplantation.

Kidney transplantation has become a well-established treatment option for patients with end-stage renal failure. The persisting organ shortage remains a ...

An Immunological Model for Heterotopic Heart and Cardiac Muscle Cell Transplantation in Rats

Heterotopic heart transplantation in rats has been a commonly used model for diverse immunological studies for more than 50 years. Several modifications have been reported since the first description in 1964. After 30 years of performing heterotopic heart transplantation in rats, we have developed a simplified surgical approach, which can be easily taught and performed without further surgical training or background.
After dissection of the ascending aorta and the pulmonary artery and ligation of superior and inferior caval and pulmonary veins, the donor heart is harvested and subsequently perfused with ice-cold saline solution supplemented with heparin. After clamping and incising the recipient abdominal vessels, the donor ascending aorta and pulmonary artery are anastomosed to the recipient abdominal aorta and inferior vena cava, respectively, using continuous running sutures.
Depending on different donor-recipient combinations, this model allows analyses of either acute or chronic rejection of allografts. The immunological significance of this model is further enhanced by a novel approach of in-ear injection of vital cardiac muscle cells and subsequent analysis of draining cervical lymphatic tissue.

We describe a model of heterotopic abdominal heart transplantation in rats, implying modifications of current strategies, which lead to a simplified surgical approach. Additionally, we describe a novel rejection model by in-ear injection of vital cardiac muscle cells, allowing further transplant immunological analyses in rats.

Heterotopic heart transplantation in rats has been a commonly used model for diverse immunological studies for more than 50 years. Several modifications ...

Immunology and Infection

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, normothermic ex vivo liver perfusion (NEVLP) has been introduced as a method to evaluate and modify organ function. NEVLP has many advantages in comparison to hypothermic and subnormothermic perfusion including reduced preservation injury, restoration of normal organ function under physiologic conditions, assessment of organ performance, and as a platform for organ repair, remodeling, and modification. Both murine and porcine NEVLP models have been described. We demonstrate a rat model of NEVLP and use this model to show one of its important applications &#8212; the use of a therapeutic molecule added to liver perfusate. Catalase is an endogenous reactive oxygen species (ROS) scavenger and has been demonstrated to decrease ischemia-reperfusion in the eye, brain, and lung. Pegylation has been shown to target catalase to the endothelium. Here, we added pegylated-catalase (PEG-CAT) to the base perfusate and demonstrated its ability to mitigate liver preservation injury. An advantage of our rodent NEVLP model is that it is inexpensive in comparison to larger animal models. A limitation of this study is that it does not currently include post-perfusion liver transplantation. Therefore, prediction of the function of the organ post-transplantation cannot be made with certainty. However, the rat liver transplant model is well established and certainly could be used in conjunction with this model. In conclusion, we have demonstrated an inexpensive, simple, easily replicable NEVLP model using rats. Applications of this model can include testing novel perfusates and perfusate additives, testing software designed for organ evaluation, and experiments designed to repair organs.

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant liver donor shortage, and criteria for liver donors have been expanded. Normothermic ex vivo liver perfusion (NEVLP) has been developed to evaluate and modify organ function. This study demonstrates a rat model of NEVLP and tests the ability of pegylated-catalase, to mitigate liver preservation injury.

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, ...

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 Porcine Heterotopic Heart Transplantation Protocol for Delivery of Therapeutics to a Cardiac Allograft

Cardiac transplantation is the gold standard treatment for end-stage heart failure. However, it remains limited by the number of available donor hearts and complications such as primary graft dysfunction and graft rejection. The recent clinical use of an ex vivo perfusion device in cardiac transplantation introduces a unique opportunity for treating cardiac allografts with therapeutic interventions to improve function and avoid deleterious recipient responses. Establishing a translational, large-animal model for therapeutic delivery to the entire allograft is essential for testing novel therapeutic approaches in cardiac transplantation. The porcine, heterotopic heart transplantation model in the intraabdominal position serves as an excellent model for assessing the effects of novel interventions and the immunopathology of graft rejection. This model additionally offers long-term survival for the pig, given that the graft is not required to maintain the recipient's circulation. The aim of this protocol is to provide a reproducible and robust approach for achieving ex vivo delivery of a therapeutic to the entire cardiac allograft prior to transplantation and provide technical details to perform a survival heterotopic transplant of the ex vivo perfused heart.

We present a protocol for utilizing a normothermic ex vivo sanguinous perfusion system for the delivery of therapeutics to an entire cardiac allograft in a porcine heterotopic heart transplant model.

Cardiac transplantation is the gold standard treatment for end-stage heart failure. However, it remains limited by the number of available donor hearts ...

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 ...

A Heterotopic Rat Heart Transplantation Model using Circulatory Death Donor Hearts

The objective of this protocol is to set up a rat heterotopic heart transplantation model with donation after circulatory death (DCD) donor hearts. There are two setups for this protocol: heart donor setup and recipient setup. In the heart donor setup, Sprague Dawley rats are anesthetized, endotracheally intubated, and ventilated. The right carotid artery is cannulated to deliver heparin and the paralytic agent vecuronium-bromide. The DCD process is initiated by terminating the ventilation. After 20 min, the heart is exposed and the aorta distal to the brachiocephalic branch is clamped. At 25 min from terminating the ventilator, ice-cold University of Wisconsin (UW) solution is perfused through the carotid catheter to flush the heart. The heart is procured by dividing the aorta, pulmonary artery, venae cavae, and pulmonary veins and stored in UW solution for implantation. In the recipient setup, the Lewis rat is anesthetized with isoflurane. Slow-release buprenorphine is administered subcutaneously to facilitate a smooth postoperative recovery. Through a midline abdominal incision, the infra-renal aorta and the inferior vena cava are isolated and clamped with an atraumatic vascular clamp. The donor heart aorta and pulmonary artery are sutured to the recipient abdominal aorta and vena cava, respectively, with a running 8-0 Prolene. The vascular clamp is removed to reperfuse the heart. The abdominal wall is closed and the rat is recovered. After a set interval (24 h to 2 weeks), the recipient rat is anesthetized, the transplanted heart is exposed, and a balloon-tip-catheter is inserted into the left ventricle via the apex to record developed pressure and dP/dt using a data acquisition system. The heart tissue is collected for histology, immunology, or molecular analysis. A successful DCD donor rat heart transplantation model will allow further studies on the cardioprotective approaches to improve heart transplantation outcomes from DCD donors.

Here, we describe the preparation and technical details of murine heterotopic heart transplantation utilizing a circulatory death donor heart.

The objective of this protocol is to set up a rat heterotopic heart transplantation model with donation after circulatory death (DCD) donor hearts. There ...

Large-Animal Model of Donation after Circulatory Death and Normothermic Regional Perfusion for Cardiac Assessment

The increase in demand for cardiac transplantation throughout the years has fueled interest in donation after circulatory death (DCD) to expand the organ donor pool. However, the DCD process is associated with the risk of cardiac tissue injury due to the inevitable period of warm ischemia. Normothermic regional perfusion (NRP) allows for an in situ organ assessment, allowing the procurement of hearts determined to be viable. Here, we describe a clinically relevant large-animal model of DCD followed by NRP. Circulatory death is established in anesthetized pigs by stopping mechanical ventilation. After a preset warm ischemia period, an extracorporeal membrane oxygenator (ECMO) is used for a NRP period lasting at least 30 min. During this reperfusion period, the model allows the collection of various myocardial biopsies and blood samples for initial cardiac evaluation. Once NRP is weaned, biochemical, hemodynamic, and echocardiographic assessments of cardiac function and metabolism can be performed before organ procurement. This protocol closely simulates the clinical scenario previously described for DCD and NRP in heart transplantation and has the potential to facilitate studies aimed at decreasing ischemia-reperfusion injury and enhance cardiac functional preservation and recovery.

The protocol describes a large-animal (porcine) model of donation after circulatory death, followed by thoracoabdominal normothermic regional perfusion that closely simulates the clinical scenario in heart transplantation, and has the potential to facilitate therapeutic studies and strategies.

The increase in demand for cardiac transplantation throughout the years has fueled interest in donation after circulatory death (DCD) to expand the organ ...

Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation

Summary

Explore More Videos

Chapters in this video

A Modified Method for Heterotopic Mouse Heart Transplantion

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

Normothermic Ex Vivo Kidney Perfusion for the Preservation of Kidney Grafts prior to Transplantation

An Immunological Model for Heterotopic Heart and Cardiac Muscle Cell Transplantation in Rats

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

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

A Porcine Heterotopic Heart Transplantation Protocol for Delivery of Therapeutics to a Cardiac Allograft

A Modified Cuff Technique for Mouse Cervical Heterotopic Heart Transplantation Model

A Heterotopic Rat Heart Transplantation Model using Circulatory Death Donor Hearts

Large-Animal Model of Donation after Circulatory Death and Normothermic Regional Perfusion for Cardiac Assessment