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This work was sponsored by the Frontier Biotechnology Key Project of the National Key R &#38; D Program of the Ministry of Science and Technology of China (2023YFC3404300), the National Key Research and Development Program of China (2023YFF0724701), the Clinical Research Funds for Central High-Level Hospitals of Fuwai Hospital, Chinese Academy of Medical Sciences, Preclinical Study on Xenogeneic Multi-Gene-Edited Donor Pig Hearts for Orthotopic Heart Transplantation in Children with Heart Failure (2025-GSP-ZD-4), and the Project of the State Key Laboratory of Cardiovascular Diseases, Fuwai Hospital, Chinese Academy of Medical Sciences, Screening and Development of Drug Targets for Long-Term Survival of Orthotopic Cardiac Xenotransplantation (2025-GSP-GG-42).

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
<ol>
	<li>Cuff preparation
	<ol>
		<li>Use polyimide tubing to fabricate the cuffs for transplantation surgery. For the arterial cuff, select polyimide tubing with an inner diameter of 0.3&#8211;0.5 mm and cut it into 2 mm segments under a stereomicroscope (10&#215;&#8211;20&#215; magnification).</li>
		<li>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&#8211;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.</li>
	</ol>
	</li>
	<li>Mouse preparation: Use neonatal Sprague-Dawley (SD) rats aged 10&#8211;12 days as donors, and C57BL/6 or Balb/c mice aged 7&#8211;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.</li>
</ol>
2. Donor heart harvesting 
<ol>
	<li>Fixation and anesthesia
	<ol>
		<li>Anesthetize neonatal rats via intraperitoneal injection of a ketamine/xylazine mixture, such as ketamine at 80&#8211;100 mg/kg and xylazine at 5&#8211;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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Heparinization
	<ol>
		<li>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.</li>
		<li>Inject heparinized saline (0.2&#8211;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.</li>
	</ol>
	</li>
	<li>Cardiac arrest
	<ol>
		<li>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.</li>
		<li>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 &#215; 1 cm pieces and pre-soak them in ice-cold saline before use.</li>
	</ol>
	</li>
	<li>Harvesting the donor heart
	<ol>
		<li>Ligate the superior and inferior vena cava with an 8&#8211;0 silk suture, then transect them. Cut the aorta at the aortic arch and sever the pulmonary artery at its bifurcation.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
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.
<ol>
	<li>Anesthesia
	<ol>
		<li>Anesthetize the recipient mouse using the same method and place it in a supine position for stabilization.</li>
	</ol>
	</li>
	<li>Disinfection
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Skin incision
	<ol>
		<li>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&#8211;0 silk suture or cauterized with an electrocautery pen to prevent bleeding.</li>
	</ol>
	</li>
	<li>External jugular vein preparation
	<ol>
		<li>After exposing the external jugular vein, individually cauterize the branches using an electrocautery pen set to low coagulation mode (5&#8211;10 W). Place two 8-0 silk sutures near the distal end of the vein for ligation.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Carotid artery preparation
	<ol>
		<li>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.</li>
		<li>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. 
		&#8203;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.</li>
	</ol>
	</li>
</ol>
4. Donor heart anastomosis 
<ol>
	<li>Aorta-to-carotid artery anastomosis
	<ol>
		<li>After trimming the donor heart, immediately place it in the recipient mouse&#39;s neck. Use an 8-0 silk suture with a loose knot to create a loop around the donor aorta.</li>
		<li>Slip the donor aorta over the arterial cuff on the recipient&#39;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&#8217;s blood.</li>
	</ol>
	</li>
	<li>Pulmonary artery-to-external jugular vein anastomosis
	<ol>
		<li>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&#8217;s external jugular vein and secure it with an 8-0 silk ligature.</li>
	</ol>
	</li>
	<li>Cardiac resumption
	<ol>
		<li>Release the vascular clamps on the recipient&#8217;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.</li>
	</ol>
	</li>
	<li>Closure of the incision
	<ol>
		<li>Once the donor heart regains a stable rhythm, adjust its position and close the skin incision on the recipient&#8217;s neck with 4-0 monofilament sutures. Disinfect the surgical site with povidone-iodine.</li>
	</ol>
	</li>
</ol>
5. Postoperative care 
<ol>
	<li>Postoperative analgesia
	<ol>
		<li>Post-operative analgesia was maintained by subcutaneous administration of meloxicam at 5 mg/kg, dissolved in sterile saline, according to the institutionally approved protocol.</li>
	</ol>
	</li>
	<li>Recovery
	<ol>
		<li>Place the mouse on a warm soft pad maintained at 37&#8211;38 &#176;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.</li>
	</ol>
	</li>
	<li>Monitoring
	<ol>
		<li>Monitor the transplanted heart daily by visual inspection and palpation to assess its beating and overall graft viability. 
		&#8203;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.</li>
	</ol>
	</li>
</ol>

<ol><li>Ali, A., Kemter, E., Wolf, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Advances+in+organ+and+tissue+xenotransplantation%5BTitle%5D)+AND+%22Annu+Rev+Anim+Biosci%22%5BJournal%5D)">Advances in organ and tissue xenotransplantation.</a> Annu Rev Anim Biosci. 12, 369-390 (2024).</li>
<li>Griffith, B. P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Genetically+modified+porcine-to-human+cardiac+xenotransplantation%5BTitle%5D)+AND+%22N+Engl+J+Med%22%5BJournal%5D)">Genetically modified porcine-to-human cardiac xenotransplantation.</a> N Engl J Med. 387 (1), 35-44 (2022).</li>
<li>Griffith, B. P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Transplantation+of+a+genetically+modified+porcine+heart+into+a+live+human%5BTitle%5D)+AND+%22Nat+Med%22%5BJournal%5D)">Transplantation of a genetically modified porcine heart into a live human.</a> Nat Med. 31 (2), 589-598 (2025).</li>
<li>Mohiuddin, M. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Graft+dysfunction+in+compassionate+use+of+genetically+engineered+pig-to-human+cardiac+xenotransplantation%3A+A+case+report%5BTitle%5D)+AND+%22Lancet%22%5BJournal%5D)">Graft dysfunction in compassionate use of genetically engineered pig-to-human cardiac xenotransplantation: A case report.</a> Lancet. 402 (10399), 397-410 (2023).</li>
<li>Kawai, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Xenotransplantation+of+a+porcine+kidney+for+end-stage+kidney+disease%5BTitle%5D)+AND+%22N+Engl+J+Med%22%5BJournal%5D)">Xenotransplantation of a porcine kidney for end-stage kidney disease.</a> N Engl J Med. 392 (19), 1933-1940 (2025).</li>
<li>Lu, T. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Advances+in+innate+immunity+to+overcome+immune+rejection+during+xenotransplantation%5BTitle%5D)+AND+%22Cells%22%5BJournal%5D)">Advances in innate immunity to overcome immune rejection during xenotransplantation.</a> Cells. 11 (23), 3865(2022).</li>
<li>Zhou, Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Current+status+of+xenotransplantation+research+and+the+strategies+for+preventing+xenograft+rejection%5BTitle%5D)+AND+%22Front+Immunol%22%5BJournal%5D)">Current status of xenotransplantation research and the strategies for preventing xenograft rejection.</a> Front Immunol. 13, 928173(2022).</li>
<li>Habibabady, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Antibody-mediated+rejection+in+xenotransplantation%3A+Can+it+be+prevented+or+reversed%5BTitle%5D)+AND+%22Xenotransplantation%22%5BJournal%5D)">Antibody-mediated rejection in xenotransplantation: Can it be prevented or reversed.</a> Xenotransplantation. 30 (4), e12816(2023).</li>
<li>Loupy, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Immune+response+after+pig-to-human+kidney+xenotransplantation%3A+a+multimodal+phenotyping+study%5BTitle%5D)+AND+%22Lancet%22%5BJournal%5D)">Immune response after pig-to-human kidney xenotransplantation: a multimodal phenotyping study.</a> Lancet. 402 (10408), 1158-1169 (2023).</li>
<li>Kinoshita, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Novel+factors+potentially+initiating+acute+antibody-mediated+rejection+in+pig+kidney+xenografts+despite+an+efficient+immunosuppressive+regimen%5BTitle%5D)+AND+%22Xenotransplantation%22%5BJournal%5D)">Novel factors potentially initiating acute antibody-mediated rejection in pig kidney xenografts despite an efficient immunosuppressive regimen.</a> Xenotransplantation. 31 (2), e12859(2024).</li>
<li>Tseng, H. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Animal+models+for+heart+transplantation+focusing+on+the+pathological+conditions%5BTitle%5D)+AND+%22Biomedicines%22%5BJournal%5D)">Animal models for heart transplantation focusing on the pathological conditions.</a> Biomedicines. 11 (5), 1414(2023).</li>
<li>von Bibra, C., Hinkel, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Non-human+primate+studies+for+cardiomyocyte+transplantation%E2%80%94ready+for+translation%5BTitle%5D)+AND+%22Front+Pharmacol%22%5BJournal%5D)">Non-human primate studies for cardiomyocyte transplantation—ready for translation.</a> Front Pharmacol. 15, 1408679(2024).</li>
<li>Jia, H., Chang, Y., Song, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+pig+as+an+optimal+animal+model+for+cardiovascular+research%5BTitle%5D)+AND+%22Lab+Anim+NY%22%5BJournal%5D)">The pig as an optimal animal model for cardiovascular research.</a> Lab Anim NY. 53 (6), 136-147 (2024).</li>
<li>Sorge, R. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Olfactory+exposure+to+males%2C+including+men%2C+causes+stress+and+related+analgesia+in+rodents%5BTitle%5D)+AND+%22Nat+Methods%22%5BJournal%5D)">Olfactory exposure to males, including men, causes stress and related analgesia in rodents.</a> Nat Methods. 11 (6), 629-632 (2014).</li>
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The authors declare no conflicts of interest.

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 outcomes14,15. 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...

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 University2,3,4,5. 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 cytokines6,7. 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 failure8,9,10. 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 results11,12,13. 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.

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

a standardized murine model with a three-person workflow for studying acute cellular and antibody-mediated rejection in xenotransplantation

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

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.

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...

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.

Despite recent advancements in xenotransplantation, acute rejection remains a major barrier to its clinical application. Acute cellular rejection (ACR) ...

Our research focuses on standardized small animal xenotransplantation models to study mechanisms of acute cellular and antibody mediated rejection. Existing models often suffer from operator vulnerability and lung procedures. This protocol provides a rapid, standardized, and reproducible workflow.This figure illustrates the standardized three-person workflow for murine xenatransplantation, in which operators simultaneously perform recipient preparation, donor heart procurement, and donor heart anastomosis. To begin, obtain 10 to 12 day old neonatal sprayed dolly rats and seven to eight week old C57 Black six or BALB/c mice. House all donor rats and recipient mice in a specific pathogen-free environment.With povidone iodine-soaked cotton balls, disinfect the thoracoabdominal area of the anesthetized rat. Incise the abdomen with scissors. Then use a cotton swab to gently move the intestines aside to expose the inferior vena cava.Use an insulin syringe to inject 0.2 to 0.5 milliliters of heparinized saline into the inferior vena cava. Gently compress the injection site with a cotton ball to prevent bleeding. Incise the diaphragm.Bilaterally cut the chest wall and retract it to fully expose the heart. Then remove the thymus and sever the superior vena cava to relieve pressure. Perfuse one milliliter of University of Wisconsin solution into the aorta to induce cardiac arrest.Continue perfusion until the heart changes from red to pale. Place a gauze pad soaked in ice-cold saline over the heart to lower its temperature and prevent dehydration. Ligate the superior and inferior vena cava with an 8.0 silk suture.Transect both vessels. Then cut the aorta at the aortic arch. Sever the pulmonary artery at its bifurcation.Ligate all pulmonary veins with the 8-O silk suture. Then excise the donor heart. Place the harvested heart in ice cold saline for temporary preservation.Using a depilatory cream, remove hair from the neck region of the anesthetized recipient mouse. Disinfect the skin with alternating rounds of povidone iodine and alcohol three times each. Make a two centimeter longitudinal incision approximately one centimeter to the right of the midline on the neck with a surgical blade.Perform blunt dissection to remove subcutaneous fat and muscle layers. Excise the submandibular gland. After exposing the external jugular vein, use an electrocautery pen set to low coagulation mode to individually cauterize the branches.Place two 8.0 silk sutures near the distal end of the vein for ligation. Then use microsurgical scissors to transect the external jugular vein between the two ligatures. Insert the external jugular vein through the venous cuff and push the cuff to the base of the vein.Then secure it with a vascular clamp. Use microsurgical scissors to remove the ligatures. Then flush the residual blood from the vein using heparinized saline.With a pair of microsurgical forceps, evert the external jugular vein over the cuff and secure it with an 8.0 silk suture. Perform blunt dissection posterior to the sternocleidomastoid muscle to locate the carotid sheath. Open the carotid sheath.Carefully separate the common carotid artery from the internal jugular vein and vagus nerve. Next, ligate and transect the carotid artery. Then insert the lower end of the transected artery through the arterial cuff.Evert the artery over the cuff and secure it in place. Slip the donor aorta over the arterial cuff on the recipient carotid artery. Secure it with the 8.0 silk ligature.Create a loose knot loop around the donor pulmonary artery. Slip the pulmonary artery over the venous cuff on the recipient external jugular vein. Then secure it with the silk ligature.Release the vascular clamps on the recipient carotid artery and external jugular vein to restore blood flow. Observe the donor heart resume strong contractions and rapidly turn red within 10 seconds. Once the donor heart regains a stable rhythm, adjust its position.Using 4.0 sutures, close the skin incision on the neck. Disinfect the surgical site with povidone iodine. Place the mouse on a warm, soft pad maintained at 37 to 38 degrees Celsius to preserve body temperature.Monitor the mouse until it regains consciousness. House the mouse individually in a cage with adequate access to food and water once fully awake. The median graft survival time in BALB/c recipients was four days.Whereas in C57 Black 6 recipients, it was significantly prolonged to 6.5 days. In the SD-to-BALB/c group, hematoxalin and eosin staining showed severe myocardial destruction with widespread erythrocyte extravasation. In the SD-to-C57 Black 6 group, hematoxalin and eosin staining showed myocardial damage characterized predominantly by inflammatory infiltrates.In the SD-to-BALB/c group, immunofluorescent staining showed minimal CD3 positive T-cell infiltration, minimal CD68 positive macrophage infiltration, and intense widespread C4D deposition. In the SD-to-C57 Black 6 group, immunofluorescent staining showed prominent CD3 positive T-cell infiltration, moderate CD68 positive macrophage presence, and moderate C4D deposition. The average success rate was 93.3%in the three-person group, compared with 86.7%in the single person group.The main challenge in team correlation, earning practice, circularization and the assure stable and fashioned modern establishment. Future studies can use this standardized model to investigate xenograft immunomechanisms and evaluate normal enumosuppressive or tolerance-inducing stretches.

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Research

JoVE Journal

Medicine

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In Vitro and In Vivo Assessment of T, B and Myeloid Cells Suppressive Activity and Humoral Responses from Transplant Recipients

This study presents a pre-clinical heterotopic abdominal heart xenotransplantation model using α-Gal knockout (GTKO) pigs and macaques. The model incorporates optimized recipient selection, vascular anastomosis techniques, and perioperative management strategies. It is suitable for evaluating novel immunosuppressive agents and provides a convenient platform for advancing heart xenotransplantation.

A Standardized Pig to Macaque Heterotopic Heart Xenotransplantation Model

Transplantation of isolated islets has been proposed to be a potential treatment for type 1 diabetes.  Here we describe a method to isolate islets from mouse pancreata and transplant them to the subcapsular space of the kidney.

A Method for Murine Islet Isolation and Subcapsular Kidney Transplantation

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

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