Cervical Heterotopic Mouse Heart Transplantation using End-to-side Anastomosis Technique to Study Transplant Immunology

Summary

This protocol presents a simplified, end-to-side suture anastomosis technique for murine cervical heart transplantation, providing a robust and reproducible model for studying transplant immunology.

Abstract

Murine heterotopic heart transplantation is a critical preclinical model in transplantation research. Cervical heterotopic heart transplantation in mice offers distinct advantages, including direct visual and palpable monitoring of graft viability, reduced surgical trauma, and faster postoperative recovery. It has therefore become an increasingly valuable platform for studying transplant immunology, particularly valuable for animals with poor physiological conditions due to genetic modification or drug induction, and for experiments requiring secondary or combined organ transplantation, the cervical region often becomes the only feasible site. Conventional cervical transplantation techniques, whether using end-to-end suturing or cuff-based methods, require permanent ligation of the recipient’s common carotid artery and external jugular vein, which may compromise cerebral perfusion and introduce foreign materials or hemodynamic disturbances. To address these limitations, this paper details a refined cervical heart transplantation technique utilizing an end-to-side vascular anastomosis. The method preserves the recipient's common carotid artery and external jugular vein, maintaining normal cerebral perfusion and avoiding complications associated with permanent vessel ligation. This protocol details the steps from donor heart harvest to recipient anastomosis and postoperative care. Representative results demonstrate a high surgical success rate (96.7%) and the model's effectiveness in studying allograft rejection. This technique offers a physiologically relevant and technically accessible model for investigators in transplant immunology.

Introduction

Over the past half‑century, transplantation immunology has continued to be a central focus for immunologists and transplant specialists. Today, solid organ transplantation is widely performed and represents the most effective treatment for end-stage organ diseases. However, graft rejection has not been fundamentally resolved and still represents one of the major obstacles in transplantation and a leading cause of graft loss¹. Consequently, overcoming or mitigating rejection while reducing or avoiding systemic immunosuppression remains a significant challenge and a key research objective in the field.

Vascularized animal organ transplantation models, particularly murine models, have played a crucial role in advancing transplant immunology and have provided an essential scientific foundation for clinical translation². For example, many immunosuppressive agents used clinically today—such as cyclosporine, tacrolimus, and rapamycin—were first successfully validated in mouse transplantation models^3,4.

The mouse heterotopic heart transplantation (HT) model is one of the most widely used preclinical models, favored due to the abundance of available genetic strains and research reagents, as well as its relatively simple procedure and easy postoperative monitoring. As such, it serves as a key platform for studying acute rejection mechanisms, inducing immune tolerance, and evaluating new immunosuppressants in vivo^5,6.

The first murine heterotopic HT model was established by Corry et al.⁷ in 1973. In this model, the donor heart is placed in the recipient's abdominal cavity, with the graft aorta and pulmonary artery (PA) anastomosed end-to-side to the recipient's abdominal aorta and inferior vena cava (IVC), respectively. This method remained the standard technique for abdominal HT for decades. Driven by the need to accommodate third-party allografts in tolerance studies, a second site for HT was explored. This led to the development of cervical HT models in mice, which offer other advantages such as direct visual assessment of graft viability, less surgical trauma, and lower risk of postoperative infection^5,8,9. In 1991, Chen et al.⁸ first described a cervical heterotopic HT model in mice, involving end-to-end manual suture anastomosis of the donor aorta and PA to the recipient's common carotid artery (CCA) and external jugular vein (EJV), respectively. However, this model did not gain widespread adoption due to the significant technical challenge posed by the severe mismatch between the donor aorta and the recipient CCA, which made vascular anastomosis extremely difficult.

Also in 1991, Matsuura et al.⁹ introduced a non-suture cuff technique for vascular anastomosis in the cervical HT model. This method simplifies the procedure by eliminating the technically demanding manual vascular suturing, making it easier to master. Subsequently, numerous technical refinements to the cuff technique have been reported^10,11. Currently, the cuff method is widely used in cervical HT, particularly by researchers without prior experience in microvascular suturing. However, the cuff technique introduces artificial materials, which may induce local inflammation and interfere with immunological assessments, while also posing technical challenges in small-caliber vessels^5,6. Recently, Liao et al.¹² proposed a cuff-free sleeve technique in cervical HT, in which donor and recipient vessels are directly anastomosed using a sleeve method. By avoiding artificial materials and accommodating small vessels, this approach offers a physiological and reproducible platform for secondary heart transplantation.

Nevertheless, both end-to-end suture and non-suture cuff techniques require permanent ligation of the recipient's unilateral CCA and EJV. This compromises unilateral cerebral blood supply to the central nervous system, which may lead to neurological complications, such as directional disorder and hemiplegia^13,14,15. Furthermore, these end-to-end approaches can lead to altered hemodynamics, such as high-velocity arterial jets, and carry a higher risk of thrombosis and stenosis, which are detrimental for long-term graft survival. Moreover, the cuff technique also carries an inherent risk of thrombosis due to the presence of the cuff.

To address these limitations, we adopted a refined end-to-side manual suture technique where the donor's ascending aorta and PA are anastomosed to the recipient's CCA and EJV, respectively. This method preserves the integrity and continuity of the recipient's major cervical vessels, thereby maintaining normal cerebral perfusion and adhering to the 3Rs (Replacement, Reduction, and Refinement) principle of animal research. Using this technique, we achieved a surgical success rate of approximately 96.7% (29/30, with only one mortality due to hemorrhage). No cerebral ischemia-related complications were observed in any recipient mice post-surgery. This method closely resembles standard abdominal models and is particularly suitable for surgeons with vascular suturing experience to establish cervical HT models.

Protocol

Animal care complied with international guidelines, and all procedures were conducted at the Institute of Organ Transplantation, Tongji Hospital, Tongji Medical College, Huazhong University of Science and Technology, China. Ethical approval for the study was obtained from the Animal Ethics Committee of Tongji Hospital (Approval number: TJH-202310010). The appropriate instruments used for the study are listed in the Table of Materials.

1. Preoperative Preparation

1. Sterilize the operating room using an ultraviolet lamp for 30 min. Disinfect the operating table with 70% ethanol and autoclave all surgical instruments (121 °C for 20 min).
2. Use adult male C57BL/6 and BALB/c mice (weighing 20–25 g; 8–10 weeks old). For syngeneic transplantation, use C57BL/6 mice as both donors and recipients. For allogeneic transplantation, use BALB/c mice as donors and C57BL/6 mice as recipients.
3. Administer preoperative analgesia to the recipient mouse via subcutaneous injection of buprenorphine (0.05 mg/kg) 1 h prior to the surgery.
   NOTE: Preoperative analgesia is not performed on the donor mouse, as the donor is euthanized immediately following the heart harvesting procedure.
4. Prepare the anesthetic solution containing ketamine (20 mg/mL) and xylazine (2 mg/mL). Anesthetize the mouse via intraperitoneal injection at 5 μL/g body weight, corresponding to 100 mg/kg ketamine and 10 mg/kg xylazine. Apply ophthalmic ointment to both eyes after induction of anesthesia to prevent corneal drying. Confirm adequate anesthesia by observing slow and regular respiration, loss of righting reflex, and absence of pedal reflex. 
5. Shave fur from the thoracic and abdominal areas. Secure the mouse in a supine position on a heating pad to maintain body temperature. Disinfect the surgical site by alternating povidone-iodine and 70% ethanol for three cycles. Extend the recipient’s neck by retracting the upper incisors upward with a 6‑0 silk thread to optimize exposure.
6. Wear sterile surgical gloves. Make an oval-shaped opening slightly larger than the incision site on a sterile drape. Then use it to cover the recipient mouse.

2. Donor heart harvest

1. Make a midline abdominal incision followed by a transverse (cruciform) incision to fully expose the peritoneal cavity.
2. Gently retract the small intestine to the left using a sterile cotton swab moistened with saline to expose the abdominal aorta and the IVC.
3. Inject 0.3 mL of heparinized saline (300 IU/mL) into the IVC and wait 1 min for systemic heparinization. Transect the abdominal aorta and IVC to exsanguinate the donor.
4. Incise the diaphragm along with the anterior chest wall to make a U-shaped incision through the rib cage using scissors to expose the thoracic cavity.
5. Inject 2 mL of cold saline (4 °C) into the heart through the intrathoracic IVC to flush out the residual blood.
   NOTE: The primary purpose of this step is to induce hypothermia and prompt the heart to stop beating rapidly. The second purpose is to thoroughly flush blood out of the heart, especially from the coronary arteries.
6. Cover the heart with a piece of sterile gauze soaked in cold saline (4 °C). Gentle downward traction on this gauze facilitates optimal exposure of the aorta and PA. Furthermore, the cold saline-soaked gauze serves to maintain hypothermia of the graft during the subsequent dissection.
7. Identify and isolate the intrathoracic IVC. Ligate it using a 6-0 silk suture and transect distal to the ligation with microscissors.
8. Identify and remove the thymus to expose the aortic arch and PA. Identify and isolate the superior vena cava. Ligate it using a 6-0 silk suture. Transect it distal to the ligation.
9. Identify and isolate the ascending aorta. Transect the aorta proximal to the innominate artery to maximize vessel length.
10. Identify and isolate the PA. Transect it just proximal to its bifurcation. Ligate all vessels posterior to the heart (the pulmonary veins) with a 6-0 silk. Carefully excise the heart using microscissors.
11. Gently move the heart and immerse it in cold saline (4 °C) placed on ice for storage until transplantation.

3. Recipient operation

1. Recipient vessel isolation
   1. Make a 1 cm longitudinal incision on the right side of the neck, extending from the superior border of the midpoint of the right clavicle to the inferior border of the mandible.
   2. Excise the right lobe of the submandibular gland. Transect the right sternocleidomastoid muscle to create a sufficient pocket for donor heart implantation.
   3. Identify and isolate the right EJV and the right CCA. Mobilize the EJV by cauterizing its branches. Then, isolate the CCA from the surrounding tissues. Avoid damaging the accompanying vagus nerve when dissecting the CCA.
2. Artery anastomosis
   1. Simultaneously occlude the EJV and CCA using two microvascular clamps, with one clamp applied to the cranial ends of both vessels and the other to the caudal ends. Apply the cranial clamp first, followed by the caudal clamp.
      NOTE: The microvascular clamps are preferably introduced from the right side of the mouse toward the left, with the clamp handles positioned on the right side. This orientation facilitates the subsequent vascular anastomosis and helps reduce compression of the trachea.
   2. Rotate the mouse 90° clockwise. Make a longitudinal incision on the CCA, ensuring the length matches the diameter of the donor ascending aorta. Flush the lumen with heparinized saline to thoroughly remove any residual blood clots.
   3. To create the anastomotic window, use an 11-0 suture. Pass the needle through the vessel wall at a distance roughly equal to the donor vessel’s diameter, gently lifting the vessel wall with the suture and then excising a small portion with micro scissors.
   4. Place the donor heart in the cervical pocket. Anchor the donor ascending aorta to the recipient CCA by placing two 11-0 sutures with needles at the proximal and distal ends. Position the donor heart on the right side of the recipient and cover it with wet sterile gauze.
   5. Using the distal suture, perform a continuous running suture on the posterior walls in a distal-to-proximal direction with 6–8 stitches.
   6. After completing the posterior wall suture, rotate the mouse 180°. Reflect the donor heart towards the recipient's left and cover it with wet sterile gauze. Complete the anastomosis by suturing the anterior vessel walls in a proximal-to-distal direction with 6–8 stitches. Flush the lumen with saline before tying the final knot to remove air bubbles.
3. Vein anastomosis
   1. Make a longitudinal incision on the EJV, matching the diameter of the donor PA. Flush the lumen with heparinized saline to thoroughly remove any residual blood clots.
   2. Anchor the donor PA to the recipient EJV by placing two 11-0 sutures with needles at the proximal and distal ends.
   3. Using the proximal suture, continuously suture the posterior walls in a proximal-to-distal direction with 5–7 stitches.
   4. Continue with the same suture to close the anterior walls from the outside in a distal-to-proximal direction with 5–7 stitches. Tie the suture at the proximal apex. Flush the lumen with saline before tying the final knot to remove air bubbles.
4. Reperfusion
   1. Release the distal clamp first, followed by the proximal clamp, to initiate reperfusion. Upon reperfusion, the cardiac graft should immediately turn red. Normal rhythm and contraction typically recover within 1–2 min.
   2. Once hemostasis is confirmed, carefully position the cardiac graft within the subcutaneous pocket, ensuring that there is no torsion of the vessels. Close the skin incision with 5-0 sutures.
5. Postoperative care
   1. Place the mouse in a clean cage and put it in an infant incubator set to 34 °C to facilitate recovery. Monitor the mouse every 1–2 h until it awakens spontaneously, then return it to the housing cage.
      NOTE: Rewarming is necessary to prevent hypothermia-induced respiratory depression and to facilitate the recovery from anesthesia.
   2. Administer postoperative analgesia via subcutaneous injection of buprenorphine (0.05 mg/kg) every 12 h for the first 48 h after surgery.
   3. Monitor the pulsations of the cardiac graft daily and record the graft survival status.

Results

Thirty cases were performed using this modified model; the operation success rate was 96.7% (29/30). Only one mortality occurred due to intraoperative hemorrhage. The remaining 29 cases were included for analysis of operative times and graft outcomes. No cerebral ischemia-related complications were observed in any recipient post-surgery. The donor heart harvest was completed in 6.4 ± 0.7 min, and the recipient operation in 28.7 ± 1.2 min. Specifically, vessel preparation, arterial anastomosis, and venous anastomosis were...

Discussion

The cervical HT model using the end-to-side anastomosis technique offers several advantages. Compared with end-to-end suturing or the cuff method, this approach completely preserves the continuity of CCA and EJV, thereby maintaining normal cerebral hemodynamics and effectively avoiding cerebral ischemia‑related complications. The end-to-side anastomosis also avoids technical difficulties caused by a mismatch in the diameters of the donor and recipient vessels. Compared with abdominal HT, this method causes less tra...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1mL Syring	Becton Dickinson Medical Devices Co., Ltd	B309628
11-0 sutures with needles	Jinhuan Medical Products Co., Ltd.	H1101	For vascular anastomosis
3mL Syring	Becton Dickinson Medical Devices Co., Ltd	B309657
5-0 sutures with needles	Jinhuan Medical Products Co., Ltd.	CR537	For skin closure
6-0 silk suture	Jinhuan Medical Products Co., Ltd.	2650182
Blood Stop Cautery	Ningbo Shunye Medical Co., Ltd.	BDD-YE-DT-1
Buprenorphine	Tianjin Institute of Pharmaceutical Research Pharmaceutical Co., Ltd	H12020275	Painkiller
Cotton swabs	Fuqing Health & Integral Medical	20230R
Curved micro forceps	RWD life Science Co.	F11031-11	Surgical tool
Curved micro scissors	66 Vision Tech Co., Ltd.	54108B	Surgical tool
Gauze swabs	Yubei Medical Materials Co., LTD	21080274
Heparin sodium injection	North China Pharmaceutical Co., LTD	2101131-2
Micro needle holder	World Precision Instruments Co., Ltd.	555408NT	Surgical tool
Micro vessel clip	Roboz Surgical Instrument Co	RS-5481T
Micro Vessel clip applicator	Roboz Surgical Instrument Co	RS-5480
Needle holder	RWD life Science Co.	F31025-13	Abdominal closure
Saline	Zhejiang Tianrui Pharmaceutical Co.,Ltd	716092103
Straight micro forceps	RWD life Science Co.	F11029-11	Surgical tool
Surgical microscope	Leica	M650
Surgical platform			Custom, magnetic
Tissue forceps	RWD life Science Co.	F12002-12	Surgical tool
Tissue scissor	RWD life Science Co.	S12003-09	Surgical tool