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

Summary

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.

Abstract

Murine heterotopic cardiac transplantation is a well-established model for investigating allograft rejection and immune tolerance. In tolerance studies, the placement of a second heterotopic cardiac graft is often required to assess donor specificity, with the cervical region providing an ideal anatomical site. While cuff-based techniques simplify vascular anastomoses, these methods introduce artificial materials that can promote peri-anastomotic inflammation, alter hemodynamics, and confound immunological analyses. Additionally, vessel narrowing resulting from genetic modifications, immunosuppressive treatment, advanced recipient age, or chronic experimental conditions can make cuff placement technically challenging in small-caliber vessels. Here, we present a cuff-free, suture-based technique for cervical heterotopic heart transplantation in mice. This method avoids the use of intraluminal polymers, preserves physiological blood flow, and reduces complications associated with altered flow dynamics. The protocol includes meticulous preparation of both the donor and recipient vessels, end-to-end micro-anastomoses of the carotid artery and external jugular vein, intraoperative patency assessments, and postoperative monitoring strategies. By minimizing the use of artificial materials and adapting to small-caliber vessels, this cuff-free technique establishes a physiological and reproducible platform for secondary heart transplantation. This approach enables rigorous mechanistic studies of donor-specific tolerance in murine cardiac transplantation models.

Introduction

Heart transplantation is a life-saving intervention for patients with end-stage heart failure¹. Recipients with cardiac allografts require lifelong immunosuppressive therapy to prevent rejection². Despite immunosuppression, long-term graft survival remains compromised by acute and chronic rejection, particularly antibody-mediated rejection (AMR) and cardiac allograft vasculopathy (CAV)³. In addition, these regimens carry significant side effects, including nephrotoxicity, metabolic syndrome, infections, and malignancies, which reduce both graft and patient survival^2,4. These challenges underscore the urgent need for immune tolerance, a state in which the recipient's immune system accepts the donor graft without requiring lifelong immunosuppression⁵. Achieving tolerance would eliminate drug-related toxicities, reduce infection and malignancy risk, prevent chronic alloimmune injury, and allow steroid withdrawal, thereby improving metabolic control and preserving graft function. While multiple tolerance strategies have shown promise in animal models, including mixed chimerism, costimulatory blockade, regulatory cell infusions, and gene-targeted approaches, clinical translation remains limited^6,7,8,9. A significant barrier is the need for reliable in vivo models that rigorously test donor-specific tolerance⁵.

Murine heterotopic heart transplantation is a powerful and widely used experimental model for studying the mechanisms of allograft rejection and tolerance^{10,11,12,13,14}. The availability of genetically defined mouse strains and transgenic models enables a rigorous dissection of immune pathways involved in graft injury and regulation^9,15,16. In tolerance studies, a second heterotopic heart transplant, either donor-matched or third-party, is often required to evaluate the donor specificity of immune regulation^13,17,18. The cervical region provides an ideal anatomical site for this second transplant, offering direct surgical access, reliable graft monitoring via palpation or imaging, and efficient retrieval of tissues for downstream immunological and histological analyses^8,19. These features make the cervical model particularly valuable for investigating mechanisms of tolerance induction and rejection across a wide range of transplantation settings.

Several cervical heterotopic heart transplantation techniques using sutures have been described over the past three decades. Early work established that the neck is a feasible site for graft implantation and direct functional monitoring using suture-based end-to-end anastomoses between the donor's vessels and the recipient's carotid and jugular system²⁰. Other groups have reported cuff-free sleeve configurations as an alternative means of securing the donor pulmonary artery or aorta to the cervical vasculature²¹. A novel end-to-side suture-based cervical model has been proposed to preserve native carotid continuity and more closely approximate the geometry of the standard abdominal end-to-side anastomosis²². Together, these approaches highlight the versatility of the cervical site and illustrate necessary trade-offs between technical complexity, hemodynamics, and the amount of foreign material at the anastomotic interface.

One widely applied vascular anastomotic technique in murine cervical heart transplantation is the intraluminal cuff technique^23,24,25,26. Cuff-based cervical techniques introduce intraluminal stents to simplify microvascular anastomoses and increase technical success rates, particularly for less experienced microsurgeons^24,25,26. While cuff-based approaches simplify the technical challenge of connecting vessels, this technique introduces foreign material into the anastomotic site. The presence of polymers may promote peri-anastomotic inflammation, alter local hemodynamics, and potentially confound immune analyses^27,28. Furthermore, narrowing of recipient vessels, whether due to genetically modified mouse strains, immunosuppressive treatment, advanced recipient age, or chronic experimental conditions, can make cuff placement technically challenging in small-caliber vessels²⁹.

To address these limitations, we developed a fully cuff-free, suture-based technique for cervical heterotopic heart transplantation. In this configuration, the recipient carotid artery is anastomosed end-to-end to the donor ascending aorta, and the recipient external jugular vein is anastomosed end-to-end to the donor pulmonary trunk, providing straightforward inflow and outflow for coronary perfusion (Figure 1A). By minimizing the use of intraluminal synthetic material, this approach avoids direct polymer-blood contact at the anastomotic site and is conceptually attractive for studies focused on local immune and inflammatory responses. The method involves meticulous preparation of both the donor and recipient vessels, as well as end-to-end micro-anastomoses of the carotid artery and the external jugular vein. Additionally, intraoperative patency assessment and postoperative monitoring are performed. In practice, this cuff-free cervical model is applied when a secondary cardiac graft is required to test donor-specific tolerance after an abdominal graft with long-term survival, particularly in transgenic or immunosuppressed mice with small-caliber vessels. However, the technique also requires advanced microsurgical expertise and access to an operating microscope; additional limitations related to carotid ligation and recipient selection are discussed below. By providing a reproducible platform, this technique enhances both the reliability of secondary transplantation models and supports mechanistic studies of donor-specific tolerance in murine systems. This protocol employs end-to-end anastomoses by connecting the recipient carotid artery to the donor ascending aorta and the recipient external jugular vein to the donor pulmonary trunk, thereby providing robust coronary perfusion of the graft (Figure 1B). End-to-end anastomoses create a straightforward inflow-outflow pattern that reliably supports coronary perfusion in this model.

Protocol

Mice were purchased from Jackson Laboratory. Both male and female mice, aged 6 - 10 weeks and weighing 20 - 30 g at the time of the first abdominal cardiac transplant, were included in this study. Donors and recipients were housed under specific pathogen-free (SPF) conditions at the rodent facility of Massachusetts General Hospital. All procedures were performed humanely in accordance with the NIH guidelines for the care and use of laboratory animals. All animal experiments were approved by and conducted under the oversight of the Institutional Animal Care and Use Committee (IACUC) at Massachusetts General Hospital (protocol number 2020N000125). The Table of Materials lists all reagents, instruments, and equipment used in this protocol. IACUC approval is required before initiating experiments. House and maintain all mice in accordance with institutional guidelines and applicable regulatory standards.

1. Preparation of animals and instruments

1. Prior to surgery, sterilize all instruments by exposing them to saturated steam at 250 °F (121 °C) under pressure for 20 min. The required microsurgical instruments are shown in Figure 2. Protect the tips of microsurgical instruments to avoid damage. Replace broken or deformed instruments immediately to maintain optimal surgical success rates.
2. Recipient preoperative care
   1. This protocol uses recipient mice previously implanted with an abdominal heterotopic heart graft^10,11,12. Confirm that the recipients are healthy, that the primary incision is fully healed and free of infection, and that the abdominal graft continues to beat. Perform this secondary cervical transplant between postoperative day (POD) 50 - 100 following the abdominal graft. Only use recipients with a functioning abdominal graft, as this is a tolerance protocol^8,17.
   2. Recipient mice have free access to food and water before the procedure. Administer extended-release buprenorphine, 3.25 mg/kg subcutaneously before surgery begins. Administer meloxicam (5–10 mg/kg) subcutaneously before surgery begins and then every 12 h for 72 h postoperatively³⁰.
3. Donor preparation
   1. Select sex- and age-matched donor mice of the appropriate strain according to the study design. For cardiac allograft immune tolerance studies, donor groups may include syngeneic (same strain as the recipient), allogeneic donor (same strain as the primary abdominal cardiac transplant donor), and allogeneic third-party (different strain from both the primary donor and the recipient).
   2. Confirm the health status and appropriate acclimation period of all donor and recipient mice before anesthesia.

2. Anesthesia

1. Anesthesia induction and support
   1. Induce anesthesia with 4% - 5% isoflurane in oxygen (1 - 2 L/min flow) using a precision vaporizer in an induction chamber. Maintain anesthesia with 1% - 2% isoflurane via a nose cone or endotracheal tube. Apply the veterinary ophthalmic lubricant to both eyes of the mouse to prevent corneal drying under anesthesia.
   2. Confirm anesthesia depth by the absence of pedal withdrawal reflex. Place the animal on a heating pad to maintain body temperature throughout the procedure.

3. Donor heart harvest

1. Donor exposure
   1. Place the donor mouse in the supine position and secure the limbs with tape. Remove hair and disinfect the skin three times using alternating applications of 10% povidone-iodine and 70% isopropyl alcohol swabs in a circular motion.
   2. Perform a midline laparotomy followed by a midline sternotomy using fine scissors to expose the abdominal and thoracic cavity. Gently mobilize the intestines to the left side with moistened gauze. Place a saline-soaked gauze beneath the intestines to maintain hydration.
2. Perfusion and donor heart harvest
   1. Prepare cold heparinized saline (125 IU/mL) and load 0.2 - 0.3 mL into a 0.5 mL syringe.
   2. Gently cannulate the inferior vena cava with a 26 - 30G needle or catheter. Slowly perfuse 2 - 3 mL of cold heparinized saline over 30 s. Wait at least 1 - 2 min to optimize heparinization.
   3. Perform a bilateral thoracotomy to expose the heart with maximum visualization. Separate the anterior thoracic cage from the dorsal side and reflect it upward, securing it with a needle or clamp.
   4. Bluntly dissect the connective tissue between the ascending aorta (AA) and pulmonary trunk (PT; Figure 3A).
   5. Ligate the inferior and superior venae cavae with 8 - 0 nylon sutures close to the heart and transect the ligated vessels distal to the ligatures. Bluntly dissect the PT as distally as possible.
   6. Ligate the left and right pulmonary veins together with a 6 - 0 nylon suture and transect the ligated vessels distal to the ligature.
   7. Excise the heart, preserving adequate length of the AA by transecting at the origin of the brachiocephalic artery and the PT. Preserve the anterior wall of the PT as long as possible. Trim the AA and PT to ensure a secure anastomosis (Figure 3B).
   8. Place the heart in a 60 mm cell culture dish filled with chilled normal saline and gently re-perfuse the donor heart with an additional 1 mL of cold heparinized saline (125 IU/mL) through the AA or PT to flush any residual blood. Avoid high-pressure injections to prevent myocardial edema or vessel injury. Trim the vessels on the chilled plate as needed.
   9. Place the heart in cold Ringer's lactate solution on ice until implantation. The donor heart is best implanted within 90 min of cold ischemia and should not be preserved for more than 120 min.

4. Recipient cervical area exposure

1. Incision and vessel isolation
   1. Induce anesthesia with 4% - 5% isoflurane in oxygen (1 - 2 L/min flow) using a precision vaporizer in an induction chamber. Maintain anesthesia with 1% - 2% isoflurane via a nose cone or endotracheal tube. Confirm adequate depth of anesthesia by the absence of pedal withdrawal reflex. Inject 0.5 mL of warm normal saline subcutaneously into the dorsal region of the recipient for fluid replenishment.
   2. Place the recipient on a heating pad to maintain body temperature. Shave the entire cervical region. Position the mouse in the supine position with the tail facing the operator. Immobilize the head, limbs, and tail with tape, ensuring a secure fixation without overstretching. Apply one drop of ophthalmic lubricant to each eye to prevent drying. Protect the area surrounding the incision with sterile surgical drapes. Disinfect the cervical skin three times using alternating applications of 10% povidone-iodine and 70% isopropyl alcohol swabs in a circular motion.
   3. Make a longitudinal skin incision with a reversed T-shape over the neck, extending from the sternum to the right mandibular angle, using a sterile scalpel blade (Supplementary Figure 1A).
   4. Bluntly mobilize the right external jugular vein (EJV) from the right medial clavicle to its major confluence. Cauterize and transect small branches of the EJV (Figure 4B).
   5. Isolate the right lobe of the submandibular gland and transect it by cauterization. Transect the sternomastoid muscle by cauterization to expose the right carotid artery (CA).
   6. Bluntly mobilize the right CA as far as possible above the carotid bifurcation (Figure 4C).
   7. Apply a small vascular clamp to the proximal portion of the CA. Place a 10 - 0 nylon ligature around the distal portion at the level of the carotid bifurcation to occlude blood flow. To reduce the risk of thrombosis and ischemia-related events, the duration of blood flow occlusion should not exceed 60 minutes.
   8. Apply a 10 - 0 nylon ligature to the proximal portion of the EJV at the level of the CA clamp. Once the EJV distends, stop bleeding from previously cauterized and transected small-branch sites. Identify the confluence of the retromandibular and posterior auricular veins as the EJV distends and apply a small vascular clamp at this confluence to occlude flow.
2. Vascular anastomosis preparation
   1. Transect the EJV and CA between the ligature and clamp using fine scissors.
   2. Flush the vessel openings with normal saline to remove blood and clots. Keep the operative field moist throughout the graft implantation procedure.

5. Graft implantation

1. Graft placement
   1. Place the donor heart graft upside down in the recipient cervical region with the PT oriented laterally and the ascending aorta (AA) oriented medially.
   2. Cover the graft with a moistened gauze, leaving the PT and AA exposed. Gauze protects and stabilizes the graft while improving visualization.
2. Blood vessel anastomosis
   1. Anastomose the donor AA end-to-end to the recipient CA using 11 - 0 nylon sutures by sleeve technique with the donor's AA superficial bites to the recipient's CA. Avoid excessive tension (Figure 4D).
   2. Anastomose the donor PT end-to-end to the recipient EJV using 11 - 0 nylon sutures by sleeve technique with the donor's PT superficial bites to the recipient's EJV. Avoid excessive tension (Figure 4E).
   3. Remove the distal EJV clamp. Remove the proximal CA clamp to restore blood flow (Figure 4F). The graft should immediately fill with blood, become red, and contract within 1 - 2 min (Supplementary Video 1). Apply a few drops of 37 °C warm normal saline to the graft to help restore contractility.
   4. Gently compress the anastomotic sites with cotton tips to achieve hemostasis.
3. Pocket closure
   1. Position the graft securely within the cervical pocket without twisting or compressing the vessels and trachea.
   2. Close the cervical incision with 6 - 0 absorbable sutures with a simple interrupted stitch (Supplementary Figure 1B). Do not cover the surgery site with a bandage; leave the area open to air.

6. Post-operative care and monitoring

1. Immediate monitoring
   1. Place the animal in a warm recovery chamber for at least 1 h. Observe for abnormal behaviors such as disorientation, tremors, or failure to ambulate.
   2. Inject 0.3 mL of warm normal saline subcutaneously for fluid replacement. Transfer the recipient to a sterilized new cage supplied with nutritional gel. House transplanted animals individually and do not return them to group housing until full recovery, as indicated by normal mobility and behavior.
   3. Administer Meloxicam (5 - 10 mg/kg, subcutaneously) every 12 h for 72 h postoperatively. The surgeon and the veterinarian examine the animals regularly during the recovery period. Remove absorbable sutures after 7-10 days or at timing according to institutional policy.
   4. Euthanize humanely if the animal exhibits > 20% body weight loss, severe swelling at the surgical site, signs of infection, lethargy, or other indications of distress, using CO₂ inhalation according to the institutional guidelines.

7. Experimental endpoints

1. Tissue collection
   1. Harvest grafts at predetermined time points for histopathology or immunological analysis. For flow cytometry and histopathology, collect tissues 2 - 3 days prior to the expected cessation of graft pulsation, based on preliminary experiments.
   2. Collect the graft, spleen, lymph nodes, and blood samples as required by the study design.
2. Survival analysis
   1. Record graft survival time, defined as the day when palpable pulsation ceases. Define long-term survival (tolerance) in secondary cervical heart transplantation as persistent donor graft pulsation beyond 50 days post-transplantation, with histopathology confirming the absence of large areas of necrosis.
   2. Plot graft survival curves using Kaplan-Meier analysis.

Results

A schematic overview of the cuff-free cervical heterotopic heart transplantation procedure is shown in Figure 1. The microscopic view of the vascular anastomoses is shown in Figure 1A, where the donor ascending aorta is anastomosed end-to-end to the recipient carotid artery and the donor pulmonary trunk is anastomosed end-to-end to the recipient external jugular vein. Figure 1B demonstrates the blood flow pattern through the graft. ...

Discussion

Murine heart transplantation models have played a critical role in advancing our understanding of transplant immunology. Since the development of the original heterotopic abdominal model, these systems have been utilized to characterize the cellular and molecular mechanisms of rejection, test novel immunosuppressive regimens, and evaluate genetic modifications in both donor and recipient strain^8,17,31. The ability to use inbred ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.5ml insulinsyringe	Covidien	8881600350	Subcutaneously injection
0.9% Sodium Chloride Injection	BD	1727170103	Intraveneous injection
10-0 nylon suture	AROSurgical Instruments	T04A10N07-13	For mice microscopic surgery
11-0 nylon suture	ETHICON	9016G	For mice microscopic surgery
1ml Syringe	BD	309659	Intraveneous injection
30G X 1/2 Needle	BD	305106	Intraveneous injection
3ml Syringe	BD	309657	Intraveneous injection
6-0 nylon suture	ETHICON	J212H	For mice microscopic surgery
60mm cell culture plate	FALCON	353002	For graft preservation
Colored Aluminum Cases	Fine Science Tools	20330-04	For mice microscopic surgery instruments disinfection
Cotton Tips	Fisherbrand	14-960-3M	Stop bleeding
Dumont #5 Forceps	Fine Science Tools	11251-20	For mice microscopic surgery
Ethiqa XR Buprofenor (CIII) VI	RX GENERICS	86084010030	Analgesia
Fine sissors	Fine Science Tools	15400-12	For mice microscopic surgery
Gauze Sponges	MEDLINE	NON25334	Protect the graft and interstine
Heat Lamp with Base	Braintree Scientfic	HL-1B	Post surgery care
Heparin Sodium	Northstar Rxllc.	72603-412-01	Intraveneous injection
Isoflurane	Baxter Anesthetic & Crit Care	1001936060	For anesthesia
Isopropyl alcohol swabs	BD	326895	Disinfect the skin before surgery
Microscope	Carl Zeiss	303294-9903	For mice microscopic surgery
Mini Trimmer	Braintree Scientfic	CLP-88	For mice hair removal
Optixcare Eye Lube Plus	AVENTIX	OPX-4252	Ophthalmic lubricant for eye protection during surgery
Pivetal Alloxate (Meloxicam)	PIVETAL	21294589	Analgesia
Portable Cauterizing Instrument Kit	McKesson Argent	42295143	For mice microscopic surgery
Ringer’s lactate buffered solution	Thermo Scientific	AAJ67572AP	For graft preservation
Small Animal Heated Pad	K&H	HM10	For mice surgery
Transpore White Surgical Tape	3M	7100115870	Fix the limb, head, and tail of mice during procedure
Vessal clamp	Fine Science Tools	00398-02	For mice microscopic surgery