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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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 systolic afterload pressure created by the host heart in the recipient circulatory system, leading to a limited donor heart performance and underestimation in the assessment18. Therefore, non-ejecting models are more favorable in heterotopic transplantation. In non-ejecting models, the donor heart is perfused but doesn&#39;t support the recipient&#39;s circulation, significantly limiting the performance assessment of the heart. Morphological and molecular evaluations, such as histologic staining and blotting analysis, can be beneficial for examining donor heart conditions when functional assessments are limited. Moreover, the metabolic markers can be evaluated using advanced technologies, such as positron emission tomography (PET) or magnetic resonance imaging (MRI)19. This model can be useful in testing the long-term effectiveness of pharmacologic and genetic interventions before implantation.
Numerous research groups have developed a normothermic ex situ preservation model, which has been successfully employed for preserving porcine hearts for up to 12 h6. However, the maintenance of large animal models can be cost-prohibitive for small laboratories, as it involves substantial expenses and requires a considerable number of trained personnel. To address this issue, we propose a less expensive and technically straightforward ex situ preservation method, which involves the use of autologous blood followed by heterotopic heart transplantation. Notably, the cost of a single experiment using our model is approximately $300. Although there is no equivalent small animal model to compare the costs, the ex situ perfusion apparatus for large animals, when used once, can cost up to $30,00016.
The presented protocol demonstrates that all the experimental procedures can be performed in a stepwise manner by a single experimenter (Figure 3). The possibility of heterotopic implantation after ex situ preservation is another advantage of this model. By cannulating the descending aorta of the donor heart for ex situ perfusion, we were able to spare the ascending part without causing any damage. Furthermore, we modified the Langendorff circuit, reducing the amount of perfusion solution required to 12 mL for effective heart perfusion. The perfusion blood was obtained from the donor rat before harvesting, allowing us to preserve the heart with its own blood and avoid any immunologic reactions during preservation.
Modifications and troubleshooting 
The ex situ perfusion circuit is recommended to maintain a mean afterload pressure within the range of 50-70 mmHg. The pressure is determined by various factors, including perfusion flow, coronary artery resistance, and perfusate viscosity20. Coronary arterial resistance is susceptible to fluctuations due to variations in temperature and pH, thus it is crucial to maintain these parameters within the normal range. The required perfusion flow varies for each experiment and is dependent on the necessary flow to maintain the desired perfusion pressure. Typically, a flow of 3-4 mL/min (equivalent to 5-6 rpm for our pump) is sufficient for a 350-450 g rat heart. The hematocrit level is a determinant of perfusate viscosity21. For our circuit, the optimal hematocrit range is 25% to 30%. Despite the utilization of the smallest experimental oxygenator, the large gas exchange surface area of 0.05 m2 for a perfusate volume of 12 mL can lead to evaporation and consequent fluid loss over time. This fluid loss can be rectified by the addition of distilled water as required. It is not recommended to add saline or ringer solution to the perfusate, as they can cause hypernatremia. The perfusate glucose concentration should be maintained at 100-150 mg/dL.
It is crucial to avoid arrhythmia during perfusion as it signifies the deterioration of one or more physiological parameters of the ex situ environment10. Tachyarrhythmia or left ventricular fibrillation are commonly associated with various factors, such as electrolytic imbalance, low hematocrit, acidosis/alkalosis, hyperthermia, and excessive afterload. On the other hand, bradyarrhythmia is mainly caused by hypothermia. Lactate and potassium are the key parameters in assessing myocardial viability. Elevated lactate levels (&#62;5 mmol/L) and hyperkalemia (&#62;5.0 mg/dL) indicate a substantial degree of myocardial damage22.
The careful monitoring of the anesthesia dosage and breathing patterns of the recipient rat is crucial during surgical procedures. Since the animals are not ventilated, continuous administration of excessive anesthesia can lead to hypoventilation and failure. The total laparotomy and extraction of abdominal organs result in significant heat loss, which can further deteriorate the recipient&#39;s condition. Therefore, the use of a temperature controller equipped with a heating pad and temperature probe is crucial to mitigate the impact of heat loss and maintain a stable body temperature.
Critical steps 
The critical stages in the surgical procedure involve the dissection of the aortic arch and MPA, aortic cannulation for ex situ perfusion, de-airing before ex situ perfusion, and de-airing before removing the clamps after implantation. These steps are highly vulnerable and are often associated with failure. However, the key to overcoming these challenges lies in identifying the appropriate technique and gaining sufficient practice. During vessel isolation in the recipient, particular attention must be paid to the right ureter, which is situated in close proximity to the IVC in the retroperitoneal space and may mimic the lymphatic duct. In the context of vein anastomosis, it is recommended to first secure the caudal end using stay sutures followed by the cranial end to prevent tearing and stenosis. This is particularly important due to the relatively fragile nature of veins in comparison to the aorta.
Limitations 
The surgical procedures involved in this experiment are considerably complex, particularly when obtaining the donor heart and perfusating blood from the same animal. The functional assessments post-implantation are limited as we utilized a non-ejecting model. An ejecting model is considered to provide more advanced outcomes in an ex situ environment. However, in heterotopic transplantation, it is constrained due to the presence of a supporting host heart in the circulatory system.

An erratum was issued for:&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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, such as lactate, potassium, and mean aortic pressure. In this study, the use of normothermic ex situ preservation decreased the total ischemic time of six successful cases to 46.2 &#177; 4.7 min, while the total out-of-body time was 166.2 &#177; 4.7 min (Figure 5). The extraction of the heart from the donor and preparation for ex situ perfusion and heterotopic transplantation required 5.8 &#177; 1.3 min, as shown in Figure 5. The overall success rate of the surgery was 70% and the mean anastomosis time of the six successful cases was 38.4 &#177; 3.4 min. In all experiments, the heart rate significantly decreased immediately after implantation, but it eventually recovered over time, as illustrated in Figure 6. The gross structure of the donor hearts was well preserved after ex situ preservation and heterotopic implantation, with no visible damages detected. However, hematoxylin-eosin staining revealed an increased number of inflammatory cells, mostly neutrophils, after 3 h of heterotopic implantation (Figure 7).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64954/64954fig01.jpg" /> 
Figure 1: Experimental design of normothermic ex situ heart preservation with heterotopic heart transplantation. Abbreviations: BGA = blood gas analysis, CPS = cardioplegic solution. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64954/64954fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64954/64954fig02.jpg" /> 
Figure 2: Schematics of modified small animal ex situ heart preservation. Abbreviations: BP sensor = blood pressure sensor, CPS = cardioplegic solution. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64954/64954fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64954/64954fig03.jpg" /> 
Figure 3: The order of anastomosis in heterotopic heart transplantation. (A) Schematics of donor heart position in the recipient abdomen and order of anastomosis. (B) Donor ascending aorta and recipient abdominal aorta anastomosis. (C) Donor pulmonary artery and recipient IVC anastomosis. Abbreviations: LV = left ventricle, RV = right ventricle, LA = left atrium, MPA = main pulmonary artery, IVC = inferior vena cava. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64954/64954fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64954/64954fig04.jpg" /> 
Figure 4: Parameters for viability assessment during ex situ perfusion. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64954/64954fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64954/64954fig05.jpg" /> 
Figure 5: Preservation timeline of the six successfully preserved hearts. Heart extraction and ex situ perfusion facilitation: 5.8 &#177; 1.3 min. Ex situ perfusion: 120 min. Implantation into the abdomen of the recipient rat: 38.4 &#177; 3.4 min. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64954/64954fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/64954/64954fig06.jpg" /> 
Figure 6: The electrophysiologic performance of the donor heart before procurement and after implantation. (A) Changes in the heart rate. Pre-harvesting, 30 min, 60 min, 90 min, 120 min, 150 min, 180 min: the times after implantation. (B) Electrocardiography images before donor heart harvesting and after 3 h of implantation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64954/64954fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/64954/64954fig07.jpg" /> 
Figure 7: Macroscopic (A-C) and microscopic (D-F) appearance of the donor heart. (A,D) Before normothermic ex situ preservation. (B,E) After normothermic ex situ preservation. (C,F) After 2 h of heterotopic implantation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64954/64954fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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 ...

rat-model-normothermic-ex-situ-perfused-heterotopic-heart

Nanyang Technological University

Research

JoVE Journal

Medicine

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

Orthotopic Aortic Transplantation: A Rat Model to Study the Development of Chronic Vasculopathy

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant vasculopathy (TVP).
The commonly used animal model for cardiovascular chronic rejection studies is the heterotopic heart transplant model performed in laboratory rodents. This model is used widely in experiments since Ono and Lindsey (3) published their technique. To analyze the findings in the blood vessels, the heart has to be sectioned and all vessels have to be measured.
Another method to investigate chronic rejection in cardiovascular questionings is the aortic transplant model (1, 2). In the orthotopic aortic transplant model, the aorta can easily be histologically evaluated (2). The PVG-to-ACI model is especially useful for CAV studies, since acute vascular rejection is not a major confounding factor and Cyclosporin A (CsA) treatment does not prevent the development of CAV, similar to what we find in the clinical setting (4). A7-day period of CsA is required in this model to prevent acute rejection and to achieve long-term survival with the development of TVP.
This model can also be used to investigate acute cellular rejection and media necrosis in xenogeneic models (5).

This video demonstrates the orthotopic aortic transplant model as a simple model to study the development of transplant vasculopathy (TVP) in rats.

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant ...

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 ...

A Novel Microsurgical Model for Heterotopic, En Bloc Chest Wall, Thymus, and Heart Transplantation in Mice

Exploration of novel strategies in organ transplantation to prolong allograft survival and minimizing the need for long-term maintenance immunosuppression must be pursued. Employing vascularized bone marrow transplantation and co-transplantation of the thymus have shown promise in this regard in various animal models.1-11 Vascularized bone marrow transplantation allows for the uninterrupted transfer of donor bone marrow cells within the preserved donor microenvironment, and the incorporation of thymus tissue with vascularized bone marrow transplantation has shown to increase T-cell chimerism ultimately playing a supportive role in the induction of immune regulation. The combination of solid organ and vascularized composite allotransplantation can uniquely combine these strategies in the form of a novel transplant model. Murine models serve as an excellent paradigm to explore the mechanisms of acute and chronic rejection, chimerism, and tolerance induction, thus providing the foundation to propagate superior allograft survival strategies for larger animal models and future clinical application. Herein, we developed a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique. The experience in syngeneic and allogeneic transplant settings is described for future broader immunological investigations via an instructional manuscript and video supplement.

To study combined solid organ and vascularized composite allotransplantation, we describe a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique.

Exploration of novel strategies in organ transplantation to prolong allograft survival and minimizing the need for long-term maintenance immunosuppression ...

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 ...

Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation

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Chapters in this video

Orthotopic Aortic Transplantation: A Rat Model to Study the Development of Chronic Vasculopathy

Murine Heterotopic Heart Transplant Technique

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

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

Ex Situ Normothermic Machine Perfusion of Donor Livers

Heterotopic Cervical Heart Transplantation in Mice

A Novel Microsurgical Model for Heterotopic, En Bloc Chest Wall, Thymus, and Heart Transplantation in Mice

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

A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents

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