Name: Cervical Heterotopic Mouse Heart Transplantation using End-to-side Anastomosis Technique to Study Transplant Immunology (Video) | JoVE Journal
Uploaded: 2026-07-03T12:30:00.000+00:00
Duration: 8 min 15 s
Description: Learn cervical heterotopic heart transplantation in mice using end-to-side anastomosis to study transplant immunology. This refined technique preserves recipient vasculature and achieves high surgical success for reliable graft monitoring.

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5 minutes

This work was supported by the National Natural Science Foundation of China (81700571).

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
<ol>
	<li>Sterilize the operating room using an ultraviolet lamp for 30 min. Disinfect the operating table with 70% ethanol and autoclave all surgical instruments (121 &#176;C for 20 min).</li>
	<li>Use adult male C57BL/6 and BALB/c mice (weighing 20&#8211;25 g; 8&#8211;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.</li>
	<li>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.</li>
	<li>Prepare the anesthetic solution containing ketamine (20 mg/mL) and xylazine (2 mg/mL). Anesthetize the mouse via intraperitoneal injection at 5 &#956;L/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.&#160;</li>
	<li>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&#8217;s neck by retracting the upper incisors upward with a 6&#8209;0 silk thread to optimize exposure.</li>
	<li>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.</li>
</ol>
2. Donor heart harvest 
<ol>
	<li>Make a midline abdominal incision followed by a transverse (cruciform) incision to fully expose the peritoneal cavity.</li>
	<li>Gently retract the small intestine to the left using a sterile cotton swab moistened with saline to expose the abdominal aorta and the IVC.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Inject 2 mL of cold saline (4 &#176;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.</li>
	<li>Cover the heart with a piece of sterile gauze soaked in cold saline (4 &#176;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.</li>
	<li>Identify and isolate the intrathoracic IVC. Ligate it using a 6-0 silk suture and transect distal to the ligation with microscissors.</li>
	<li>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.</li>
	<li>Identify and isolate the ascending aorta. Transect the aorta proximal to the innominate artery to maximize vessel length.</li>
	<li>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.</li>
	<li>Gently move the heart and immerse it in cold saline (4 &#176;C) placed on ice for storage until transplantation.</li>
</ol>
3. Recipient operation
<ol>
	<li>Recipient vessel isolation
	<ol>
		<li>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.</li>
		<li>Excise the right lobe of the submandibular gland. Transect the right sternocleidomastoid muscle to create a sufficient pocket for donor heart implantation.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Artery anastomosis
	<ol>
		<li>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.</li>
		<li>Rotate the mouse 90&#176; 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.</li>
		<li>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&#8217;s diameter, gently lifting the vessel wall with the suture and then excising a small portion with micro scissors.</li>
		<li>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.</li>
		<li>Using the distal suture, perform a continuous running suture on the posterior walls in a distal-to-proximal direction with 6&#8211;8 stitches.</li>
		<li>After completing the posterior wall suture, rotate the mouse 180&#176;. Reflect the donor heart towards the recipient&#39;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&#8211;8 stitches. Flush the lumen with saline before tying the final knot to remove air bubbles.</li>
	</ol>
	</li>
	<li>Vein anastomosis
	<ol>
		<li>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.</li>
		<li>Anchor the donor PA to the recipient EJV by placing two 11-0 sutures with needles at the proximal and distal ends.</li>
		<li>Using the proximal suture, continuously suture the posterior walls in a proximal-to-distal direction with 5&#8211;7 stitches.</li>
		<li>Continue with the same suture to close the anterior walls from the outside in a distal-to-proximal direction with 5&#8211;7 stitches. Tie the suture at the proximal apex. Flush the lumen with saline before tying the final knot to remove air bubbles.</li>
	</ol>
	</li>
	<li>Reperfusion
	<ol>
		<li>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&#8211;2 min.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Postoperative care
	<ol>
		<li>Place the mouse in a clean cage and put it in an infant incubator set to 34 &#176;C to facilitate recovery. Monitor the mouse every 1&#8211;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.</li>
		<li>Administer postoperative analgesia via subcutaneous injection of buprenorphine (0.05 mg/kg) every 12 h for the first 48 h after surgery.</li>
		<li>Monitor the pulsations of the cardiac graft daily and record the graft survival status.</li>
	</ol>
	</li>
</ol>

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The authors have no conflicts of interest to declare.

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&#8209;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...

Over the past half&#8209;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&#8212;such as cyclosporine, tacrolimus, and rapamycin&#8212;were first successfully validated in mouse transplantation models3,4.
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 vivo5,6.
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&#39;s abdominal cavity, with the graft aorta and pulmonary artery (PA) anastomosed end-to-side to the recipient&#39;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 infection5,8,9. 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&#39;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 reported10,11. 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 vessels5,6. 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&#39;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 hemiplegia13,14,15. 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&#39;s ascending aorta and PA are anastomosed to the recipient&#39;s CCA and EJV, respectively. This method preserves the integrity and continuity of the recipient&#39;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.

<table><tbody><tr><td>1mL Syring</td><td>Becton Dickinson Medical Devices Co., Ltd</td><td>B309628</td><td></td></tr><tr><td>11-0 sutures with needles</td><td>Jinhuan Medical Products Co., Ltd.</td><td>H1101</td><td>For vascular anastomosis</td></tr><tr><td>3mL Syring</td><td>Becton Dickinson Medical Devices Co., Ltd</td><td>B309657</td><td></td></tr><tr><td>5-0 sutures with needles</td><td>Jinhuan Medical Products Co., Ltd.</td><td>CR537</td><td>For skin closure</td></tr><tr><td>6-0 silk suture</td><td>Jinhuan Medical Products Co., Ltd.</td><td>2650182</td><td></td></tr><tr><td>Blood Stop Cautery</td><td>&lt;a name=&quot;RANGE!B7&quot;&gt;Ningbo Shunye Medical Co., Ltd.&lt;/a&gt;</td><td>BDD-YE-DT-1</td><td></td></tr><tr><td>Buprenorphine</td><td>Tianjin Institute of Pharmaceutical Research Pharmaceutical Co., Ltd</td><td>H12020275</td><td>Painkiller</td></tr><tr><td>Cotton swabs</td><td>Fuqing Health &amp;amp; Integral Medical</td><td>20230R</td><td></td></tr><tr><td>Curved micro forceps</td><td>RWD life Science Co.</td><td>F11031-11</td><td>Surgical tool</td></tr><tr><td>Curved micro scissors</td><td>66 Vision Tech Co., Ltd.</td><td>54108B</td><td>Surgical tool</td></tr><tr><td>Gauze swabs</td><td>Yubei Medical Materials Co., LTD</td><td>21080274</td><td></td></tr><tr><td>Heparin sodium injection</td><td>North China Pharmaceutical Co., LTD</td><td>2101131-2</td><td></td></tr><tr><td>Micro needle holder</td><td>World Precision Instruments Co., Ltd.</td><td>555408NT</td><td>Surgical tool</td></tr><tr><td>Micro vessel clip</td><td>Roboz Surgical Instrument Co</td><td>RS-5481T</td><td></td></tr><tr><td>Micro Vessel clip applicator</td><td>Roboz Surgical Instrument Co</td><td>RS-5480</td><td></td></tr><tr><td>Needle holder</td><td>RWD life Science Co.</td><td>F31025-13</td><td>Abdominal closure</td></tr><tr><td>Saline</td><td>Zhejiang Tianrui Pharmaceutical Co.,Ltd</td><td>716092103</td><td></td></tr><tr><td>Straight micro forceps</td><td>RWD life Science Co.</td><td>F11029-11</td><td>Surgical tool</td></tr><tr><td>Surgical microscope</td><td>Leica</td><td>M650</td><td></td></tr><tr><td>Surgical platform</td><td></td><td></td><td>Custom, magnetic</td></tr><tr><td>Tissue forceps</td><td>RWD life Science Co.</td><td>F12002-12</td><td>Surgical tool</td></tr><tr><td>Tissue scissor</td><td>RWD life Science Co.</td><td>S12003-09</td><td>Surgical tool</td></tr></tbody></table>

cervical heterotopic mouse heart transplantation using end-to-side anastomosis technique to study transplant immunology

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

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&#8217;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&#39;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&#39;s effectiveness in studying allograft rejection. This technique offers a physiologically relevant and technically accessible model for investigators in transplant immunology.

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...

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.

Murine heterotopic heart transplantation is a critical preclinical model in transplantation research. Cervical heterotopic heart transplantation in mice ...

We study transplant immunology using refined mice cervical heart transplant model with end-to-side anastomosis. Compared to traditional cuff method, our end-to-side anastomosis avoids foreign material, preserves green indocyanine, and improves long-term graft survival. To begin, expose the peritoneal cavity of the anesthetized donor mouse covered with a sterile surgical gauze.After retracting the small intestine, expose the abdominal aorta and the inferior vena cava. Inject 0.3 milliliters of heparinized saline into the inferior vena cava, and wait for one minute to allow systemic heparinization. Use scissors to transect the abdominal aorta and inferior vena cava to exsanguinate the donor.Use scissors to incise the diaphragm, along with the anterior chest wall, to create a U-shaped incision through the ribcage and expose the thoracic cavity. Inject two milliliters of cold saline at four degrees Celsius into the heart through the intrathoracic inferior vena cava to flush residual blood. Cover the heart with crushed ice and gauze soaked in ice-cold saline, and keep the gauze moist with additional ice-cold saline to maintain hypothermia.Identify and isolate the intrathoracic inferior vena cava. Ligate it using a 6-0 silk suture. Using microscissors, transect the vessel distal to the ligation.Identify and remove the thymus to expose the aortic arch and pulmonary artery. Identify and isolate the superior vena cava. Ligate it using a 6-0 silk suture, and transect distal to the ligation.Next, identify and isolate the ascending aorta. Transect the aorta proximal to the innominate artery to maximize vessel length. Identify and isolate the pulmonary artery.Transect it just proximal to its bifurcation. Ligate all vessels posterior to the heart, including the pulmonary veins, with a 6-0 silk suture. Use microscissors to carefully excise the heart.Remove the attached lung tissue and any excess surrounding tissue. Gently transfer the heart into cold saline placed on ice for storage until transplantation. Through the longitudinal neck incision on the anesthetized recipient mouse, carefully excise the right submandibular gland using forceps and a microcautery pen.Next, mobilize the external jugular vein by cauterizing its branches. Transect the right sternocleidomastoid muscle to create a sufficient pocket for donor heart implantation. Isolate the common carotid artery while avoiding damage to the vagus nerve.Use two microvascular clamps to first clamp the cranial ends of the vessels, then the caudal ends, to simultaneously occlude the external jugular vein and common carotid artery for anastomosis and reduce tracheal compression. After rotating the mouse 90 degrees clockwise, use an 11-0 suture to pass the needle through the vessel wall, and lift it at a distance roughly equal to the donor vessel's diameter. Use microscissors to excise a small portion of the vessel wall to create an anastomotic window.Flush the lumen with heparinized saline to remove residual blood clots. Rotate the mouse again to place the donor heart in the cervical pocket. Anchor the donor ascending aorta to the recipient common carotid artery using two 11-0 sutures at proximal and distal ends.Position the heart on the right side and cover it with wet sterile gauze. Using the distal suture, perform a continuous running suture on the posterior walls in a distal-to-proximal direction with six to eight stitches. Rotate the mouse 180 degrees.Reflect the donor heart to the left and cover it with wet sterile gauze. Flush the lumen with saline. Perform the anterior wall anastomosis in a proximal-to-distal direction with six to eight stitches.Make a longitudinal incision on the external jugular vein matching the diameter of the donor pulmonary artery. Flush the lumen with heparinized saline to thoroughly remove residual blood. Anchor the donor pulmonary artery to the recipient external jugular vein using two 11-0 sutures at proximal and distal ends.Using the proximal suture, continuously suture the posterior walls in a proximal-to-distal direction with five to seven stitches. Continue suturing the anterior walls from outside in a distal-to-proximal direction, and after flushing the lumen with saline, tie at the proximal apex. After covering the sutures with sterile cotton, rotate the mouse and release the distal clamp first, followed by the proximal clamp, to initiate reperfusion.Observe the cardiac graft turning red and regaining normal rhythm and contraction within one to two minutes. Confirm hemostasis and position the cardiac graft within the subcutaneous pocket. Close the skin incision using 5-0 sutures.Survival analysis of 10 syngeneic and 10 allogenic recipients showed that recipients of syngeneic grafts survived for over 100 days with normal contraction rhythm and strength. In contrast, recipients with allogeneic grafts showed typical rejection within six to eight days post-transplantation. Histological evaluation revealed that syngeneic grafts maintained intact myocardial architecture without inflammation or graft vasculopathy at postoperative days seven and 100.Conversely, allogeneic grafts at postoperative day seven exhibited severe acute cellular rejection, marked my massive lymphocyte infiltration, cardiomyocyte edema, and necrosis. This protocol is particularly suitable for genetically or drug-induced FRET mice and secondary or combined transplantations. Manual suture of tiny cervical vessels is technically challenging.This method's so certain with clear abdominal heart transplant appearance. Future studies can explore tolerance mechanisms, secondary transplantation, combined organ transplantation models, and novel immunosuppressive strategies.

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Research

JoVE Journal

Medicine

The mouse heterotopic heart transplantation model has been proven by many investigators to be an important method for studying mechanisms of rejection and immune response.  However, the techniques involved are still challenging.  By modifying standard techniques we have had success with more than 1000 transplants.

Heterotopic Heart Transplantation in Mice

The murine cervical heart transplantation model is well suited for immunological as well as ischemia reperfusion injury studies. We modified the procedure using a non-suture cuff technique and performed more than 1,000 successful transplants with this approach. Herein, we provide additional details of this technique to supplement the video.

Murine Cervical Heart Transplantation Model Using a Modified Cuff Technique

A technique is demonstrated for the microsurgical procedure for heterotopic transplantation of hearts in mice, including simplified methods for donor harvesting and recipient vessel anastomosis.

A Modified Method for Heterotopic Mouse Heart Transplantion

The manuscript describes the steps required to perform the heterotopic heart transplant in the mouse.

Murine Heterotopic Heart Transplant Technique

The cuff technique shortens and facilitates the model of heterotopic cervical heart transplantation in mice by avoiding technically challenging suture anastomoses of small vessels. The technical details shown in this video paper should allow researches to establish this model in their laboratories.

Heterotopic Cervical Heart Transplantation in Mice

To study combined solid organ and vascularized composite allotransplantation, we describe a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique.

A Novel Microsurgical Model for Heterotopic, En Bloc Chest Wall, Thymus, and Heart Transplantation in Mice

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Blood Circuit Reconstruction in an Abdominal Mouse Heart Transplantation Model

In the present protocol, a mouse heart transplantation model is used for investigating the mechanism of cardiac allograft rejection. In this heterotopic heart transplantation model, operation efficiency is improved, and the survival of cardiac grafts is ensured by a cervical end-to-end anastomosis of heart implantation using a modified Cuff technique.

A Modified Cuff Technique for Mouse Cervical Heterotopic Heart Transplantation Model

Mouse cardiac transplantation models represent valuable research tools for studying transplantation immunology. The present protocol details mouse heterotopic cervical cardiac transplantation that involves the placement of cuffs on the recipient's common carotid artery and the donor's pulmonary artery trunk to allow for laminar blood flow.

Mouse Heterotopic Cervical Cardiac Transplantation Utilizing Vascular Cuffs

This protocol presents a cuff-free, suture-based cervical heterotopic heart transplantation technique in mice, designed to enable the reliable placement of a second heterotopic cardiac graft for evaluating donor-specific immune tolerance under physiological conditions with minimal interference from foreign material.

Cuff-Free Suture-Based Cervical Model for Secondary Heterotopic Heart Transplantation to Evaluate Donor-Specific Immune Tolerance in Mice

Learn cervical heterotopic heart transplantation in mice using end-to-side anastomosis to study transplant immunology. This refined technique preserves recipient vasculature and achieves high surgical success for reliable graft monitoring.

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

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