A Standardized Murine Model with a Three-Person Workflow for Studying Acute Cellular and Antibody-Mediated Rejection in Xenotransplantation

Summary

This study establishes two standardized murine xenotransplantation models to specifically replicate acute cellular rejection and acute antibody-mediated rejection. These models serve as a reliable platform for investigating mechanisms of xenograft immune rejection, evaluating immunosuppressive strategies, and advancing preclinical xenotransplantation research.

Abstract

Despite recent advancements in xenotransplantation, acute rejection remains a major barrier to its clinical application. Acute cellular rejection (ACR) and acute antibody-mediated rejection (AMR) are the primary immune responses leading to early graft failure in xenograft recipients. While large animal models such as non-human primates and genetically modified pigs have provided valuable insights, their use is limited by high costs, ethical constraints, and significant experimental variability. Small-animal models offer a practical, reproducible alternative for mechanistic studies of immune rejection. In this study, two standardized murine xenotransplantation models were developed to specifically mimic ACR and AMR. To ensure reproducibility and minimize operator-related variability, a structured three-person surgical protocol with an assembly-line workflow was implemented, enabling consistent model generation with high efficiency and quality. The resulting models not only replicate the key immunopathological features of clinical xenograft rejection but also offer a robust platform for investigating immune mechanisms and evaluating targeted immunosuppressive strategies. These models will help accelerate the preclinical development of xenotransplantation therapies.

Introduction

Xenotransplantation has emerged as one of the most promising strategies to address the shortage of donor organs for transplantation. In recent years, rapid advancements in genetic editing technologies, immune regulation strategies, and biomedical engineering have led to significant progress in this field¹. Over the past three years, multiple research teams worldwide have conducted clinical trials involving xenotransplantation in humans, including two cases of genetically modified pig heart transplantation at the University of Maryland, a pig kidney transplantation trial at Massachusetts General Hospital, and a thymokidney transplantation trial at New York University^2,3,4,5. Despite these unprecedented advancements, immune rejection remains the primary challenge limiting the clinical application of xenotransplantation. Even with highly optimized genetically modified donor organs and combined immunosuppressive regimens, rejection remains a major obstacle. Current studies indicate that acute rejection is a critical hurdle in xenotransplantation, primarily classified into acute cellular rejection (ACR) and acute antibody-mediated rejection (AMR). ACR is primarily mediated by T cells, characterized by extensive T-cell infiltration into the graft, leading to tissue damage and functional impairment. Its underlying mechanisms involve xenogeneic antigen presentation, T-cell activation, and the release of inflammatory cytokines^6,7. In contrast, AMR is driven by preformed or newly generated donor-specific antibodies. These antibodies bind to endothelial cells within the graft, triggering complement activation and a robust immune response, ultimately resulting in microvascular injury, thrombosis, and graft failure^8,9,10. Therefore, a deeper understanding of these rejection mechanisms and the development of more precise immunoregulatory strategies are essential for advancing xenotransplantation from preclinical research to long-term clinical application.

To address the challenge of acute rejection in xenotransplantation, it is essential to return to fundamental research and preclinical studies to elucidate the underlying immunological mechanisms. Large animal models, particularly non-human primates (NHPs) and genetically modified pigs, have been widely used in xenotransplantation research due to their close anatomical and physiological similarities to humans. These models provide critical insights into immune rejection mechanisms and immunomodulatory strategies. However, their application is significantly constrained by high costs, ethical considerations, and the complexity of long-term postoperative care. Moreover, the substantial variability in immune responses among individual NHPs affects the reproducibility of experimental results^11,12,13. While large animal models remain indispensable for clinical translation, their limitations in experimental feasibility and consistency underscore the urgent need for more cost-effective, scalable, and mechanistically relevant small animal models.

Given these challenges, small animal models have gained increasing importance in xenotransplantation research due to their low cost, short experimental cycles, and ease of large-scale application. These models not only provide an essential platform for investigating immune rejection mechanisms but also serve as a critical bridge between in vitro studies and large animal experiments, facilitating a more precise exploration of immunoregulatory strategies. In this study, two standardized murine xenotransplantation models that specifically mimic ACR and AMR were established. To ensure reproducibility and consistency, a standardized three-person surgical protocol and a streamlined, assembly-line approach to model generation were implemented. This standardized workflow minimizes operator-dependent variability, resulting in highly uniform heart transplantation models. These refinements provide a reliable platform for mechanistic investigations and the evaluation of novel immunosuppressive strategies. This protocol is particularly suitable for researchers conducting small-animal studies on xenotransplantation immune mechanisms and on the evaluation of immunosuppressive strategies. It offers advantages such as low cost, high reproducibility, and a standardized workflow, providing a reliable experimental foundation for subsequent large-animal and preclinical studies.

Protocol

This protocol can be performed by a single operator or by a three-person team, with each member responsible for donor heart harvesting, recipient preparation, and donor heart anastomosis, respectively. The entire procedure is completed within 10 min, enabling a rapid, high-throughput, and standardized model generation.

The animal experiments in this study were approved by the Institutional Animal Care and Use Committee (IACUC) of Fuwai Hospital, Chinese Academy of Medical Sciences (Approval No. 0108-1-70-ZX(X)-41). All procedures were performed in strict compliance with animal welfare regulations and ethical guidelines to ensure adherence to relevant standards. The reagents and the equipment used are listed in the Table of Materials.

1. Preoperative preparation

1. Cuff preparation
   1. Use polyimide tubing to fabricate the cuffs for transplantation surgery. For the arterial cuff, select polyimide tubing with an inner diameter of 0.3–0.5 mm and cut it into 2 mm segments under a stereomicroscope (10×–20× magnification).
   2. Partially cut the clamp section of each segment circumferentially in half to form a handle-like extension. For the venous cuff, select polyimide tubing with an inner diameter of 0.4–0.7 mm and cut it into 3 mm segments. Similarly, partially cut the clamp section circumferentially in half to create a handle-like extension.
      NOTE: Soak the cuffs in 75% ethanol for 30 min, then rinse thoroughly with normal saline.
2. Mouse preparation: Use neonatal Sprague-Dawley (SD) rats aged 10–12 days as donors, and C57BL/6 or Balb/c mice aged 7–8 weeks as recipients. To ensure proper care and minimize stress, transport neonatal donor rats together with the mother rat. House all animals in a specific pathogen-free (SPF) environment.

2. Donor heart harvesting

1. Fixation and anesthesia
   1. Anesthetize neonatal rats via intraperitoneal injection of a ketamine/xylazine mixture, such as ketamine at 80–100 mg/kg and xylazine at 5–10 mg/kg, following institutionally approved protocols. Provide pre-operative analgesia by subcutaneous administration of meloxicam at 5 mg/kg using sterile saline as the vehicle. Confirm adequate anesthesia by the absence of reflex responses before proceeding with the surgical procedure.
   2. Once fully anesthetized, place the rat in a supine position and secure it under a stereomicroscope. Remove hair from the thoracoabdominal surgical area and perform aseptic preparation using three alternating scrubs with povidone-iodine or chlorhexidine-based scrub followed by alcohol. Use sterile gloves, sterile instruments, and a sterile drape to maintain aseptic conditions during donor heart harvesting.
      NOTE: Handle tribromoethanol with gloves and eye protection in a ventilated area; avoid skin contact and inhalation.
2. Heparinization
   1. Disinfect the thoracoabdominal area using povidone-iodine-soaked cotton balls. Gently move the intestines aside using a cotton swab to expose the inferior vena cava.
   2. Inject heparinized saline (0.2–0.5 mL) into the inferior vena cava using an insulin syringe, followed by gentle compression of the injection site with a cotton ball to prevent bleeding.
      NOTE: Wear gloves and eye protection when preparing or injecting heparinized saline; dispose of syringes as sharps waste. Prepare heparinized saline by dissolving one vial of medical-grade sodium heparin (125 mg) in 100 mL of normal saline.
3. Cardiac arrest
   1. Incise the diaphragm, cut the chest wall bilaterally, and retract it to fully expose the heart. Remove the thymus. Sever the superior vena cava to relieve pressure, and perfuse 1 mL of University of Wisconsin (UW) solution into the aorta to induce cardiac arrest.
   2. Continue perfusion until the heart changes from red to pale. Simultaneously, place a gauze pad soaked in ice-cold saline over the heart to lower its temperature and prevent dehydration.
      NOTE: Cut sterile gauze into 1 cm × 1 cm pieces and pre-soak them in ice-cold saline before use.
4. Harvesting the donor heart
   1. Ligate the superior and inferior vena cava with an 8–0 silk suture, then transect them. Cut the aorta at the aortic arch and sever the pulmonary artery at its bifurcation.
   2. Finally, ligate all pulmonary veins with an 8-0 silk suture and excise the donor heart. Temporarily store the harvested heart in ice-cold saline for preservation.

3. Recipient preparation

NOTE: All surgeries were performed using sterile surgical gloves and sterilized microsurgical instruments. Between animals or procedures, the instruments were re-sterilized by autoclaving or rapid sterilization using a glass bead sterilizer before reuse. The surgeon changed gloves regularly according to aseptic technique requirements and immediately replaced gloves if they became contaminated or contacted non-sterile areas.

1. Anesthesia
   1. Anesthetize the recipient mouse using the same method and place it in a supine position for stabilization.
2. Disinfection
   1. Remove hair from the neck region using a depilatory cream. Then perform aseptic skin preparation using three alternating scrubs with povidone-iodine or chlorhexidine-based scrub followed by alcohol. Place a sterile drape over the surgical site to maintain aseptic conditions and keep the suture material and donor heart sterile during the procedure.
3. Skin incision
   1. Make a 2 cm longitudinal incision approximately 1 cm to the right of the midline on the neck using ophthalmic scissors. Perform blunt dissection to gradually remove the subcutaneous fat and muscle layers, and excise the submandibular gland.
      NOTE: When excising the submandibular gland, the basal blood vessels should be ligated with an 8–0 silk suture or cauterized with an electrocautery pen to prevent bleeding.
4. External jugular vein preparation
   1. After exposing the external jugular vein, individually cauterize the branches using an electrocautery pen set to low coagulation mode (5–10 W). Place two 8-0 silk sutures near the distal end of the vein for ligation.
   2. Use microsurgical scissors to transect the external jugular vein between the two ligatures. Insert the external jugular vein through the venous cuff, push the cuff to the base of the vein, and secure it with a vascular clamp.
   3. Remove the ligatures using microsurgical scissors, and flush out residual blood within the vein using heparinized saline. Evert the external jugular vein over the cuff using microsurgical forceps and fix it with an 8-0 silk suture.
      NOTE: Operate the electrocautery pen carefully and take care to avoid burns. The external jugular vein has multiple tributaries that should be carefully isolated and sealed to minimize bleeding.
5. Carotid artery preparation
   1. Perform blunt dissection posterior to the sternocleidomastoid muscle to locate the carotid sheath. Open the carotid sheath, and carefully separate the common carotid artery from the internal jugular vein and vagus nerve.
   2. Ligate and transect the artery in the same manner as the vein. Insert the lower end of the transected artery through the arterial cuff, evert it, and secure it in place.
      ​NOTE: The carotid artery, internal jugular vein, and vagus nerve are enclosed within the carotid sheath. When opening the sheath, care should be taken to avoid damaging the vagus nerve. The carotid artery has a smaller diameter than the external jugular vein but is more elastic, making eversion more challenging than that of the vein.

4. Donor heart anastomosis

1. Aorta-to-carotid artery anastomosis
   1. After trimming the donor heart, immediately place it in the recipient mouse's neck. Use an 8-0 silk suture with a loose knot to create a loop around the donor aorta.
   2. Slip the donor aorta over the arterial cuff on the recipient's carotid artery and secure it with an 8-0 silk ligature.
      NOTE: When securing the aorta, the suture should be tied above the carotid artery ligation line to prevent direct contact between the ligation site and the recipient’s blood.
2. Pulmonary artery-to-external jugular vein anastomosis
   1. Use an 8-0 silk suture with a loose knot to create a loop around the donor pulmonary artery. Slip the pulmonary artery over the venous cuff on the recipient’s external jugular vein and secure it with an 8-0 silk ligature.
3. Cardiac resumption
   1. Release the vascular clamps on the recipient’s carotid artery and external jugular vein to restore blood flow. Observe that the donor heart resumes strong contractions and rapidly turns red within 10 s.
4. Closure of the incision
   1. Once the donor heart regains a stable rhythm, adjust its position and close the skin incision on the recipient’s neck with 4-0 monofilament sutures. Disinfect the surgical site with povidone-iodine.

5. Postoperative care

1. Postoperative analgesia
   1. Post-operative analgesia was maintained by subcutaneous administration of meloxicam at 5 mg/kg, dissolved in sterile saline, according to the institutionally approved protocol.
2. Recovery
   1. Place the mouse on a warm soft pad maintained at 37–38 °C to preserve body temperature and monitor it until it regains consciousness. Once fully awake, house the mouse individually in a cage with adequate access to food and water.
3. Monitoring
   1. Monitor the transplanted heart daily by visual inspection and palpation to assess its beating and overall graft viability.
      ​NOTE: Perform all procedures using appropriate PPE and follow institutional biosafety and animal-care regulations. All chemical residues, perfusates, and animal tissues generated during the experiment should be collected and disposed of according to institutional regulations. Solutions containing tribromoethanol or heparin should be treated as hazardous chemical waste, while animal tissues and blood-derived materials should be handled as biological waste and sterilized by autoclaving or disposed of through certified facilities.

Results

Figure 1 demonstrates the standardized three-person collaborative workflow for generating a murine xenotransplantation model. In this system, three operators work in parallel, each performing a designated task repeatedly: recipient preparation, donor heart procurement, or donor heart anastomosis. This division of labor allows one complete transplant model to be finished every 10 min. While a single operator can complete one model in approximately 30 min, the three-person workflow enables the...

Discussion

Murine xenotransplantation is a highly invasive procedure that demands a high level of technical proficiency and consistency from the operator. During model establishment, the operator has a significant impact on experimental outcomes^14,15. In a single-person workflow, the complexity and length of the procedure result in an approximate operation time of 30 min per model, leading to low efficiency. Additionally, individual differences in anatomical dissection, vas...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Anti-CD3 antibody	Abcam	ab16669
Anti-CD68 antibody	Bioss	bs-1432R
DAPI	Thermofisher	P36971
Depilatory cream	Veet	-
Electrocautery pen	Yuyan Instrument	-
Fluorescent secondary antibodies	Thermofisher	A-11008
HE staining kit	Beyotime	C0105S
Ice maker	Shjingmi	IMS-20
Insulin syringes	KDL	U-40
Lidocaine gel	Tide Pharm
Meloxicam	Sigma-Aldrich	444800
Microsurgical forceps	ZHENHAO	FS217
Microsurgical needle holders	BIOFIVEN	HH01231
Microsurgical scissors	ZHENHAO	FS237
Normal saline	Kelun Pharmaceutical	-
Polyimide tubing	Guangzhou Wanlixin Metal Products Co. , Ltd.	-	Inner diameter: 0.3-0.5 mm
Polyimide tubing	Guangzhou Wanlixin Metal Products Co. , Ltd.	-	Inner diameter: 0.4-0.7 mm
Povidone-iodine solution	INOHV	-
Sodium heparin	Qianhong bio-pharma	-
Stereomicroscope	Murzider	MSD204
Surgical sutures	Jinhuan Medical	8-0 and 4-0 silk sutures
Surgical warming pad	Hani Pet	-
Tribromoethanol	Jitianbio	JT0781
University of Wisconsin solution	Belzer	-
Vascular clamps	RWD Life	R31005-10