Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation

Summary

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

Abstract

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.

Introduction

Heart transplantation remains the sole viable therapy for refractory heart failure^1,2,3,4. 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 donors^6,7,8,9.

Normothermic ex situ heart perfusion (NESHP) using organ care system (OCS) machines has emerged as a clinical intervention^1,3. This technique has been deemed a suitable alternative to the conventional static cold storage (SCS) method^2,9. NESHP effectively reduces the duration of cold ischemia, diminishes metabolic demand, and facilitates optimal nutritional supply and oxygenation during the transportation of donor organs^10,11. 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 technique^12,13. 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.

Protocol

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.

1. 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.
2. 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.
3. Place the temperature sensor within the reservoir where the donor heart will be stored, maintaining the circuit's temperature at 37 °C.
4. Surgical preparations
   1. Prepare a separate set of sterile micro-instruments and materials for each donor and recipient rat.
      1. 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.
      2. 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.

2. Donor heart preservation and blood collection

1. Induce anesthesia in the donor rat with isoflurane (5%) in the anesthesia chamber and record the rat's weight before placing it on the surgical table.
2. 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.
3. 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.
4. 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.
5. Catheterization
   1. 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.
   2. 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.
   3. 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.
   4. 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.
6. Blood collection
   1. Collect around 10 mL of blood via the catheter inserted in the Abd. A.
   2. 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).
7. Cardiac arrest
   1. 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.
   2. 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.
   3. Remove the thymus entirely using micro forceps to visualize the aortic arch. Apply light compression if thymic arteries bleed.
8. Extraction
   1. After administering all the CPS, isolate the aortic arch from the surrounding tissues. Carefully dissect just below the left subclavian artery.
   2. 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.
   3. 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.
   4. 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.

3. Ex situ perfusion

1. Aorta cannulation and perfusion
   1. Before aorta cannulation, replace the saline-primed circuit with blood priming.
   2. 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.
   3. Confirm the correct position of the cannula by gently grasping the aorta with micro forceps.
   4. 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.
   5. Monitor the perfusion pressure and temperature through the sensor connected to the monitoring system.
   6. Gently massage the heart with the first and index fingers until venous blood leaks from the main pulmonary artery (MPA).
   7. Secure the aorta with a 1-0 silk ligature and remove the clamp after verifying all the settings (perfusion circuit, perfusion pressure, temperature).
   8. 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 °C indicates adequate perfusion.
   9. 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.

4. Implantation

1. Preparation of recipient
   1. Begin the recipient preparation 30 min before the cessation of ex situ perfusion.
   2. Anesthetize the recipient animal using the same method as mentioned in step 2.2.
   3. 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 °C.
   4. Apply eye lubricant, shave the pubic to the epigastric area, and cleanse the area with an iodine-based scrub and 70% alcohol.
2. Medications
   1. Inject 2 mL of warm saline subcutaneously to compensate for the fluid lost during the surgery. Inject 200 IU of heparin subcutaneously.
   2. Administer antibiotic prophylaxis by injecting 10 mg/kg cefazolin dissolved in 0.3 mL of saline subcutaneously or intramuscularly.
   3. Administer pain control by injecting 20 mg/kg of diclofenac subcutaneously.
3. 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.
4. Prevent dehydration by wrapping the abdominal organs with warm and wet gauze. Intermittingly spread warm saline with a 50 mL syringe during the surgery.
5. 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.
   1. Assuming vascular anastomosis to be placed infrarenal, prepare a sufficient portion of the aorta and IVC for clamping.
   2. Perform blunt preparation using cotton swabs or sharp-serrated forceps to remove the fats and fascia around the vessels.
   3. 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.
   4. 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.
   5. 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.
   6. 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.
   7. Starting the anastomoses, placed the donor heart on the right side of the recipient's abdomen and attach the donor Asc. A to the recipient's Abd. A with one simple interrupted stitch (9-0 polypropylene) at the cranial corner of the longitudinal incision.
   8. Move the heart to the left side of the recipient abdomen and perform anastomosis of the donor's Asc. A with the recipient's Abd. A using a running 9-0 polypropylene suture.
   9. 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.
   10. 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.
6. De-airing and de-clamping
   1. Remove the mesenteric vein clamp first after completing the anastomosis to allow the right side of the heart to fill with venous blood.
   2. Remove the air in the coronary circuit and Asc. A. by applying retrograde coronary perfusion for several seconds.
   3. Place a piece of gauze on both sides of the vessels and remove the caudal clamp and the cranial clamp.
   4. 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's body temperature is below 35 °C, the heart rhythm will normalize after the temperature reaches 36 °C.
7. Replace the abdominal organs in a meander-like manner and close the layers of the abdominal incision using continuous 5-0 polypropylene sutures.
8. After the surgery, place the anesthetized animal on a clean area over a heating pad until the body temperature reaches 37°C. 
   NOTE: Do not initiate the postoperative examinations till the body temperature reaches 37°C. Maintain anesthesia at 2-2.5% isoflurane until the end of the experiments.
9. 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.

Results

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, ...

Discussion

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...

Materials

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