A Standardized Pig to Macaque Heterotopic Heart Xenotransplantation Model

Heart xenotransplantation offers new hope for patients with end-stage heart failure who require a transplant but are unable to receive a suitable donor organ due to donor shortage. In pre-clinical evaluation studies, life-supporting heart xenotransplantation has achieved survival durations of up to nine months, meeting the efficacy criteria required for initiating human clinical trials¹. Based on these findings, the United States Food and Drug Administration (FDA) approved expanded access use for two clinical cases of heart xenotransplantation using genetically edited donor hearts. Although these clinical cases have demonstrated the feasibility of heart xenotransplantation, several unresolved challenges have emerged, including abnormal cardiac hypertrophy and antibody-mediated rejection^2,3,4,5. Therefore, further pre-clinical studies in large animal models under good laboratory practice conditions are necessary to provide additional data and to develop novel immunosuppressive regimens to ensure the safety of future human clinical trials.

The long-term survival of orthotopic heart xenografts is not solely limited by rejection, but also by primary graft dysfunction. The overall incidence of primary dysfunction has been reported to range from approximately 40% to 60% across different research teams^4,6,7. Due to the heightened susceptibility of porcine donor hearts to ischemic injury, some groups have developed hypothermic perfusion devices to reduce the incidence of primary graft dysfunction. However, postoperative cardiac hypertrophy remains difficult to control. Even with growth hormone receptor (GHR) knockout donor pigs, cardiac function may be compromised by the absence of GHR, thereby limiting long-term graft viability^2,3. In this context, heterotopic heart xenotransplantation presents several advantages: it avoids excessive preload and afterload, simplifies the surgical procedure, and reduces total ischemic time^8,9. This model better reflects graft injury driven by xenogeneic rejection, and unlike orthotopic transplantation, heterotopic heart xenotransplantation enables long-term graft survival, repeated laparotomies for in vivo biopsies, and prolonged observation of immune activation and dynamic immune cell function. It serves as a valuable model for investigating xenogeneic immune rejection mechanisms and for evaluating the efficacy and safety of new immunosuppressive strategies^10,11.

However, most abdominal heterotopic heart xenotransplantation models have utilized baboons as recipients, but the procurement of baboons as experimental animals remains challenging in many countries. In contrast, macaques are the most widely used non-human primate and have high phylogenetic homology with humans¹², making them a practical alternative for xenotransplantation research. Nevertheless, their significantly smaller body size compared to baboons necessitates specific surgical protocol adaptations to accommodate their distinct anatomical dimensions.

This study presents a step-by-step protocol for GTKO pig to macaque heterotopic heart xenotransplantation. The use of α-Gal knockout pigs (GTKO) is primarily aimed at preventing hyperacute rejection, which would otherwise lead to graft failure within minutes after heart xenotransplantation. It is structured as follows: (1) matching principles between donor pigs and recipient macaques; (2) optimization of the surgical procedure for heterotopic heart xenotransplantation; and (3) perioperative management. This study aims to disseminate these transplant techniques to facilitate comparability across research teams and enhance the reproducibility and translational relevance of heart xenotransplantation studies.

This study was approved by the Animal Ethics Committee at Fuwai Hospital (FWAEC-JL-010-1/0-2020). Male macaques weighing >9 kg and aged 10-15 years, and male Bama miniature pigs weighing 5-6 kg (~60 days old) were used to minimize abdominal compression. Larger-bodied macaques were prioritized to prevent abdominal compression by the donor heart and to accommodate post-transplant growth. The reagents and equipment used are listed in the Table of Materials.

1. Donor and recipient selection criteria

1. Recipient macaque selection
   1. Pre-existing antibody screening: Use flow cytometry to evaluate the level of preformed xenogeneic antibodies, enabling the selection of recipients with low immunological risk.
      NOTE: In this assay, GTKO porcine endothelial cells from GTKO pig aortas are isolated through enzymatic digestion, as described previously¹³.
      1. Incubate GTKO porcine endothelial cells with recipient plasma.
      2. Quantify binding of macaque IgM and IgG to porcine cells using fluorescently labeled secondary antibodies against macaque IgM and IgG. Select recipients with low preformed antibody levels.
   2. Baseline health evaluation: Perform serum biochemistry, complete blood count, and coagulation function tests to confirm adequate baseline organ function.
2. Donor pig selection criteria
   1. Echocardiographic screening: Select donor pigs with normal left and right ventricular function, no structural or functional abnormalities of the cardiac valves, an ejection fraction greater than 65%, and no evidence of pulmonary hypertension.
   2. Sex matching: Select donor pigs with the same sex as the recipient.
   3. Blood type: Select donor pigs with blood type O.

2. Preoperative preparation

1. Anesthesia and sedation
   1. Anesthesia: Induce macaques with ketamine (10 mg/kg) and midazolam (1 mg/kg) (following institutional approved protocol). Following induction, maintain anesthesia with isoflurane (2%-3%) plus dexmedetomidine (1 µg/kg/h) and propofol (1-2 mg/kg/h).
   2. Intubation: Intubate the donor pig with a 5.0-mm endotracheal tube. Intubate the recipient macaque with a 5.5-mm endotracheal tube.
   3. Ureteral catheter insertion: Insert a 6-F double-lumen urinary catheter in the macaque; omit in donor pig.
2. Prepare and disinfect surgical fields
   1. Skin preparation: Clip hair from the chest, neck, abdomen, back, and limbs.
   2. Disinfection: Scrub shaved areas with povidone-iodine, including hooves (pigs) or hands and feet (monkeys).
3. Central venous catheter placement in the recipient
   1. Positioning: Position the recipient macaque in a partial left lateral decubitus with shoulder elevation to optimize exposure of the right cervical region. Select the right internal jugular vein as the preferred site for central venous access.
   2. Sterilization and draping: Disinfect and drape the surgical field with povidone-iodine following standard aseptic procedures.
   3. Incision and vessel exposure: Make a 5-cm incision approximately 1 cm medial to the sternocleidomastoid muscle, parallel to its inner border. Perform a blunt dissection of the fascia and retract the muscle laterally.
      1. Identify the external jugular vein, which lies superficial and outside the carotid sheath. Carefully ligate small communicating branches of the internal jugular vein. Open the carotid sheath to expose the internal jugular vein, vagus nerve, and carotid artery.
   4. Catheter insertion: Insert an 8-F triple-lumen central venous catheter using a retrograde tunneling technique. Connect the proximal end of the catheter to the tunneling needle under sterile conditions and pull it through the tunnel into the neck incision.
      1. Exteriorize the distal end at the midpoint of the scapula. Trim the catheter to an appropriate length to ensure its tip reaches the junction of the superior vena cava and the right atrium.
   5. Incision closure: Close the wound in layers. Close the fascia and subcutaneous tissue using 3-0 absorbable sutures, and close the skin with 1-0 silk sutures. Ensure complete coverage and secure fixation of the catheter beneath the fascial layer to minimize infection risk.
      1. Use the central venous catheter as a critical fluid access pathway for postoperative medication administration and volume resuscitation in macaques. Perform daily povidone-iodine antisepsis and sterile gauze dressing to preserve catheter integrity and prevent infection.
      2. Maintain the central venous catheter in situ until the final sacrifice procedure to ensure reliable vascular access.

3. Recipient surgery (pre-implantation phase)

1. Midline abdominal incision
   1. Incision selection: Make a midline abdominal incision approximately 12-15 cm in length (Figure 1A).
   2. Incision details: Incise sequentially through the skin, superficial fascia, linea alba, transversalis fascia, and subperitoneal fascia (including extraperitoneal fat), then incise the parietal peritoneum to access the abdominal cavity. Use a #23 surgical blade for the skin incision and electrocautery for deeper tissue dissection to achieve simultaneous cutting and hemostasis.
2. Abdominal exploration and bowel management
   1. Adhesiolysis: Explore the abdominal cavity to assess for adhesions, focusing on the presence of adhesions between the intestines and the abdominal wall. If present, carefully dissect adhesions using electrocautery to minimize tissue injury and reduce the risk of postoperative mechanical bowel obstruction.
   2. Bowel protection: Gently exteriorize the small intestine and place it on sterile moist gauze for protection to expose the abdominal aorta and inferior vena cava.
3. Exposure of the abdominal aorta and inferior vena cava
   1. Mobilize the abdominal aorta and inferior vena cava: Carefully dissect surrounding tissues from the abdominal aorta and inferior vena cava, minimizing the risk of vascular injury and bleeding. Expose the abdominal aorta and inferior vena cava between the origins of the renal arteries and the bifurcation of the iliac arteries as fully as possible.
   2. Handle vascular branches: Preserve the inferior mesenteric artery and its accompanying vein during dissection. Meticulously preserve all lumbar branches.

4. Donor heart procurement

1. Median sternotomy and vessel mobilization
   1. Median sternotomy: Perform a midline sternotomy to access the thoracic cavity (Figure 1B). Carefully divide the donor sternum using a Lebsche sternum knife.
   2. Vessel mobilization: After exposing the thoracic cavity, incise the pericardium and meticulously dissect to separate the ascending aorta and main pulmonary artery.
2. Vessel clamping and cardiac cooling
   1. Vessel clamping: Place a 4-0 purse-string suture at the aortic root, then insert a single-use antegrade cardioplegia cannula for cardioplegia (Figure 1C). Sequentially clamp the superior and inferior vena cava, followed by cross-clamping the ascending aorta.
      1. Immediately after aortic cross-clamping, incise the pulmonary veins and inferior vena cava for decompression, ensuring incisions are large enough for sufficient drainage.
   2. Cardiac cooling: Suction the pericardial cavity to evacuate residual blood, then immediately immerse the heart in sterile ice-cold saline to achieve a hypothermic environment (0-4 °C).
3. Cardioplegia and perfusion monitoring
   1. Cardioplegia: Maintain perfusion pressure at 150 mmHg using a pressure bag. Perfuse the donor heart with 30 mL/kg body weight of HTK solution (e.g., 150 mL for a 5 kg pig).
   2. Perfusion monitoring: Simultaneously palpate the aortic root and left ventricle to verify proper cardioplegia delivery pressure and prevent ventricular distension. After completing cardioplegia infusion, confirm cardiac arrest by palpation and visual inspection. Ensure the absence of any myocardial fibrillation, then proceed with donor heart excision.
4. Cardiac resection and preservation
   1. Cardiac resection: After cardioplegia, apply additional ice to the pericardial cavity (Figure 1D). Gently elevate the heart and sequentially transect the inferior vena cava and pulmonary veins using surgical scissors. Take special care to avoid injury to the superior vena cava and right atrium when dividing the right superior pulmonary vein.
      1. Protect the trachea while dissociating the posterior wall of the left atrium. Once dissection reaches the level of the left atrial roof and pulmonary arteries, temporarily place the heart back into the pericardial cavity.
      2. Dissociate and transect the proximal aortic arch and superior vena cava to expose the pulmonary trunk. Continue dissection along tissue planes toward the left atrial roof, allowing removal of the heart and its great vessels.
         NOTE: Securely ligate the azygos vein stump after cardiac resection to prevent bleeding.
   2. Cardiac preservation: Immediately immerse the explanted heart in ice-cold saline for preservation. Additionally, perfuse the donor heart with another University of Wisconsin (UW) solution at a volume of 30 mL/kg, as the UW solution provides superior myocardial protection (Figure 1E).
5. Key differences from human donor heart procurement
   1. Infusion pressure settings: Set the perfusion pressure in the pressure bag to 150 mmHg, which is considered optimal for myocardial perfusion.
   2. Aortic cross-clamp position: Since pigs have only two major branches from the aortic arch, the left and right brachiocephalic trunks, place the cross-clamp between these branches. Ligate the right brachiocephalic trunk with silk sutures prior to cross-clamping to preserve a longer length of the donor aorta.
   3. Ventricular decompression strategy: To minimize ventricular overdistension and limit ischemic time, decompress only the left ventricle routinely. If the right ventricle begins to swell, open the superior vena cava above the azygos vein. If the left ventricle swells, incise the right pulmonary vein and extend the cut toward the right pulmonary hilum.

5. Donor heart implantation

1. Donor heart preparation
   1. Donor heart preparation: Securely close the donor superior vena cava, inferior vena cava, and pulmonary vein orifices of the left atrium using 4-0 prolene sutures.
   2. Preservation of the sinoatrial node: Preserve the donor superior vena cava and the sinoatrial node located near the right atrial appendage.
2. End-to-side anastomosis of the pulmonary artery to the inferior vena cava
   1. Heparinization: Administer 5,000 units of heparin sodium intravenously to the recipient macaque prior to donor heart anastomosis to prevent thrombus formation.
   2. Pulmonary artery anastomosis: Place a DeBakey atraumatic tangential clamp on the recipient inferior vena cava. Create a longitudinal incision slightly larger than the donor pulmonary artery to minimize postoperative stenosis.
      1. Using a double-armed 5-0 prolene suture, begin the anastomosis at the 12 o'clock position by inserting one needle from the inside (luminal side) of the recipient vessel outward. Insert the other needle from the donor pulmonary artery side, passing from the lumen to the outside at the corresponding 12 o'clock position.
      2. Tie the initial knot on the external vessel wall. Reverse the needle direction, passing from the outside back into the lumen of the donor pulmonary artery. Continue continuous suturing clockwise toward the 6 o'clock position, using a nerve hook to maintain suture tension.
      3. Rotate the heart to expose the opposite side, switch to the other needle, and suture the contralateral wall from 12 to 6 o'clock with a needle entry pattern of outside-to-inside on the recipient vessel and inside-to-outside on the donor vessel, completing the end-to-side anastomosis.
   3. Air evacuation: Before securing the final knot, inject saline into the anastomotic site to ensure complete air evacuation. Occlude the donor pulmonary artery using an atraumatic vascular clamp, then sequentially release the inferior vena cava vascular clamp (Figure 1F).
3. End-to-side anastomosis of the donor aorta to the recipient abdominal aorta
   1. Aortic anastomosis: Partially clamp the recipient abdominal aorta using a DeBakey atraumatic tangential clamp. Make a longitudinal incision slightly larger than the diameter of the donor aorta to prevent postoperative stenosis.
      1. Using a double-armed 5-0 prolene suture, begin the anastomosis by inserting the needle at the 12 o'clock position from the inside (luminal side) of the recipient aorta outward. Insert the other needle from the donor aorta at the 12 o'clock position, also from inside to outside, and tie the first stitch on the external vessel wall.
      2. Reverse the needle direction, passing from outside to inside on the donor vessel, and suture continuously clockwise to the 6 o'clock position, maintaining suture tension with a nerve hook. Flip the heart to expose the opposite side, reverse the needle direction accordingly (outside to inside on the donor vessel, inside to outside on the recipient vessel), and suture continuously from 12 to 6 o'clock, completing the end-to-side anastomosis.
   2. Air evacuation: Before tying the final knot, inject saline into the anastomosis to remove air. Sequentially release the abdominal aortic vascular clamp to restore circulation (Figure 1G).
4. Reperfusion and defibrillation
   1. Prevention of ventricular fibrillation: Administer 3 mg/kg lidocaine intravenously prior to reperfusion to prevent ventricular fibrillation. Observe the donor heart for spontaneous contractions within 2 min.
   2. Defibrillation: If ventricular fibrillation occurs, apply direct defibrillation by placing the paddles in direct contact with both ventricular surfaces (left and right) and deliver a biphasic 5-Joule shock. The total ischemic time of the donor heart was approximately 57.5 min ± 6.1 min (Table 1).
5. Drain placement
   1. Negative pressure drainage: Place a negative pressure drainage tube in the abdominal cavity to prevent postoperative fluid accumulation and monitor for active hemorrhage.
   2. Indication for drain removal: Monitor postoperative drainage volume and remove the abdominal drain when 24-h output falls below 25 mL.
6. Abdominal closure
   1. Abdominal closure and ultrasound examination: Close the abdominal wall in layers, approximating the fascia and subcutaneous tissue with 3-0 absorbable sutures, and close the skin with 1-0 silk sutures. Perform postoperative abdominal ultrasound to assess cardiac activity (Figure 1H).

6. Postoperative management and graft monitoring

1. Monitoring and general care guidelines
   1. Behavioral and symptom monitoring: Continuously monitor the recipient's status during the first two postoperative weeks using video surveillance. Perform daily assessments of behavior, appetite, abdominal distension, dyspnea, edema, and normal defecation and urination.
      1. Monitor body weight, general physical condition, surgical incision healing, potential central venous catheter infection, mucosal appearance, and intra-abdominal fluid accumulation.
   2. Ultrasound evaluation: Use ultrasound to evaluate the function of the heterotopically transplanted heart and detect any surrounding effusion if abnormalities are present. Figure 2 presents echocardiographic images of the xenograft during diastole and systole on postoperative day (POD) 1, alongside graft failure observed in Experiment M3. The red line indicates marked hypertrophy of the left ventricular wall in the failing xenograft.

Based on the procedural steps illustrated in Figure 1, a reproducible pig-to-macaque heterotopic heart xenotransplantation model was successfully established. The surgical process included a midline laparotomy in the recipient macaque to access the abdominal cavity and a median sternotomy in the donor pig for heart procurement. After pericardiotomy, the donor heart was arrested using HTK solution and further preserved with UW solution under topical hypothermia. Vascular anastomoses were performed in an end-to-side manner between the donor pulmonary artery and the recipient inferior vena cava, and between the donor aorta and the recipient abdominal aorta. Postoperative echocardiographic evaluation confirmed that the donor heart resumed contractile activity with preserved systolic and diastolic function, indicating successful reperfusion and viability of the graft. These results demonstrate the technical feasibility of the model and its potential for long-term functional monitoring of xenografts in non-human primates.

Figure 1: Pig-to-macaque heterotopic heart transplant: surgical steps and postoperative ultrasound. (A) The macaque's abdominal cavity was entered through a midline laparotomy with layered dissection. (B) A median sternotomy was performed to access the porcine donor's thoracic cavity, followed by pericardiotomy. (C) A perfusion cannula was inserted into the aortic root of the donor heart. (D) Following cardiac arrest induced by HTK solution perfusion, the donor heart was topically cooled with saline ice slush. (E) Subsequent perfusion with University of Wisconsin (UW) preservation solution was performed. (F) End-to-side anastomosis was created between the porcine pulmonary artery and the recipient's inferior vena cava. (G) End-to-side anastomosis was constructed between the porcine aorta and the recipient's abdominal aorta. (H) Postoperative echocardiography was conducted to assess the donor heart's systolic and diastolic function. Please click here to view a larger version of this figure.

Figure 2: Time-series echocardiographic images of M3. (A) Echocardiographic images of the xenograft during diastole and systole at postoperative day (POD) 1. (B) Echocardiographic images of the xenograft during diastole and systole at POD 3. (C) Echocardiographic images of the xenograft during diastole and systole at POD 10. (D) Echocardiographic images of the xenograft during diastole and systole at graft failure. The red line indicates a significant increase in left ventricular wall thickness during diastole. Please click here to view a larger version of this figure.

Table 1: Total ischemic time of the donor heart.

Although heterotopic abdominal heart transplantation is technically less demanding than orthotopic thoracic heart transplantation, several critical challenges must still be addressed to ensure success, particularly in small-body-weight recipients such as macaques.

First, donor-recipient immunological matching is essential, as the level of preformed antibodies in the recipient largely determines the severity of acute antibody-mediated rejection¹⁴. The current strategy involves incubating endothelial cells from GTKO pigs with plasma from potential recipient macaques, followed by fluorescence-based detection of bound IgG and IgM using anti-monkey secondary antibodies. Macaques with low levels of preformed anti-pig IgG and IgM are selected to minimize the risk of acute antibody-mediated rejection. In addition, careful control of donor heart size is important to prevent excessive intra-abdominal pressure or intestinal obstruction caused by space-occupying effects.

Attention must be paid to the presence of the left azygos vein in pigs, which drains intercostal venous blood into the coronary sinus¹⁵. Failure to identify and ligate this vein during donor preparation can lead to significant bleeding upon aortic opening, and it should be managed appropriately before procurement.

The sequence of vascular anastomosis is crucial. The procedure should begin with an end-to-side anastomosis between the donor pulmonary artery and the recipient inferior vena cava. After completing the anastomosis, the vascular clamp on the inferior vena cava can be released. Assuming preserved donor pulmonary valve function, venous blood from the recipient will not backflow into the right ventricle. As a precautionary measure, an atraumatic vascular clamp could be applied to the donor pulmonary artery to prevent retrograde blood flow and mitigate thrombotic risks. This is followed by an end-to-side anastomosis between the donor aorta and the recipient abdominal aorta. After de-airing, the aortic clamp can be released. This sequence minimizes total clamping time and maintenance of distal circulation in recipient macaques.

The donor aorta and pulmonary artery should be trimmed into an oblique oval shape rather than a vertical round shape. This configuration facilitates a more obtuse angle between the donor vessels and recipient vasculature, reducing the risk of vascular kinking or compression due to positional changes in the monkey postoperatively and ensuring smooth blood flow to the donor heart.

Perioperative anticoagulation management is critical. Continuous administration of low molecular weight heparin postoperatively is recommended to maintain the activated clotting time (ACT) at approximately 100 seconds, thereby reducing the risk of thrombosis.

This model has inherent limitations. The small body size of recipient macaques restricts the donor heart size. Beyond immunologically mediated xenograft injury and hypertrophy, the intrinsic growth potential of the porcine heart further increases the risk of abdominal organ compression, which requires consideration. Additionally, the limited blood volume of small-bodied macaques necessitates meticulous intraoperative bleeding control to reduce the risk of postoperative anemia and hypoproteinemia. An often-overlooked intraoperative challenge is the anatomical difference between pig and human hearts, specifically the presence of the left azygos vein draining into the coronary sinus in pigs. Failure to ligate this vein before reperfusion can cause severe bleeding. Attempting to repair the bleeding after the graft resumes beating is technically difficult and increases the risk of unnecessary injury to the donor heart.

In summary, compared to orthotopic heart xenotransplantation, the heterotopic model is less affected by surgical complexity or primary graft dysfunction, making it a platform for investigating xenogeneic immune rejection. It also serves as an efficient platform to evaluate genetically engineered pigs and novel immunosuppressants in vivo. The model's technical simplicity and lack of requirement for cardiopulmonary bypass enable rapid training and widespread adoption, facilitating accelerated xenotransplantation research.