A Modified Cuff Technique for Mouse Cervical Heterotopic Heart Transplantation Model

Summary

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.

Abstract

Cardiac allograft rejection limits the long-term survival of patients after heart transplantation. A mouse heart transplantation model is ideal for investigating the mechanism of cardiac allograft rejection in preclinical studies because of their high homology with human genes. This understanding would help develop unique approaches to improving patients' long-term survival treated with cardiac allografts. In a mouse model, abdominal donor heart implantation is commonly performed with an end-to-end anastomosis to the recipient's aorta and inferior vena cava using stitches. In this model, the donor's heart is implanted by end-to-end anastomosis to the recipient's carotid artery and jugular vein by the modified-Cuff technique. The transplantation surgery is performed without stitching and thus may increase the survival of the recipient since there is no interference with the blood supply and venous reflux of the lower body. This mouse model would help investigate the mechanisms underlying the immunological and pathological (acute/chronic) rejection of cardiac allografts.

Introduction

Heart transplantation has become the standard treatment for terminal heart failure. More than 5,500 heart transplantations per year are performed in the organizations registered under the International Society for Heart and Lung Transplantation. Among the allogeneic heart transplant recipients, the 1-year rejection rate is still >10%, while the 3-year rejection rate increased to 36%^1,2. However, effective prophylactic treatments for patients with cardiac allograft rejection are lacking. Therefore, animal model studies are warranted that elucidate the physiological mechanisms underlying the immunological and pathological rejection of cardiac allografts. Such studies would contribute to the investigation of novel targets required to develop efficacious drugs, which would help prevent cardiac allograft rejection and improve survival rates in those patient populations.

Some potential immunological and pathophysiological mechanisms of cardiac allograft rejection have been proposed recently in mouse model studies of heterotopic heart transplantation^3,4,5. Consequently, mouse heterotopic heart transplantation became an ideal preclinical model to investigate the mechanisms of immune rejection and pathological injury occurring in cardiac allografts after heart transplantation because of their high homology with human genes. The prevalent concept is to perform heterotopic transplantation in a mouse model by an abdominal end-to-end anastomosis in the recipient aorta and inferior vena cava using stitches, similar to the normal human anatomy. However, this procedure may interfere with the recipient's blood supply and venous reflux of the lower body⁶. Therefore, a modified heterotopic heart transplantation procedure in a mouse model is proposed here.

The donor's heart is implanted with the recipient's carotid artery and jugular vein by an end-to-end cervical anastomosis using a modified Cuff technique. This modified procedure facilitated the operative feasibility and ensured the survival of the cardiac graft without interfering with the blood supply and venous reflux of the lower body.

Protocol

All Animal experiments were performed in compliance with the Guide for the Care and Use of Laboratory Animals, Eight Edition, National Research Council (US) 2011. Procedures involving animal subjects have been approved by the Animal Care and Use Committee of the Chongqing University Cancer Hospital, Chongqing, China. Male BALB/c and C57BL/6 mice weighing 20-30 g, obtained from commercial sources (see Table of Materials), were used for allogeneic heart transplantation study. The C57BL/6 mice were used as donors and syngeneic recipients, while the BALB/c mice served as allogeneic recipients. A schematic of the protocol is shown in Figure 1.
NOTE: All supplies used during surgery, including surgical instruments and solutions, are sterile. The surgical procedure adheres to the principle of aseptic operation technique.

1. Recipient procedure

1. Induce general anaesthesia via inhalation of 5% isoflurane through a 15 x 10 x 10 cm induction chamber connected with a hood (see Table of Materials).
2. Fix the recipient mouse on the operating table with a heating pad. Maintain anaesthesia with continuous inhalation of 2% isoflurane through a face mask over the nose and mouth.
   ​NOTE: Slow respiratory rate and rhythm, the disappearance of the corneal reflex, and the absence of the pedal reflex in the toes indicate the effectiveness of anaesthesia.
3. After shaving the hair, disinfect the surgical area with three alternating rounds of povidone-iodine scrub followed by alcohol. Then, incise the skin 1.5-2 cm parallel to the cervical midline from the right mandibular angle to the tail-end.
4. Dissect ~1 cm of the right external jugular vein using an electro-coagulator and micro-forceps. Clip the vein at the proximal end with an atraumatic microvascular clamp and ligate it at the distal end.
5. Pass the distal end of the vein through a 22 G polyurethane barbed cuff (see Table of Materials) with a bevel end and superficial grooves. Fix the vein with the handle of the cuff using a microvascular clamp.
6. Remove the 8-0 ligation suture at the distal end, turn the lumen over the cuff hooked by the superficial barb inside out and fix with a 10-0 surgical suture in the grooves of the surface.
7. Resect the right sublingual gland to form a fossa for implanting the cardiac graft, and reserve the right lobe of the submaxillary gland and the right sternocleidomastoid.
8. Dissect the right common carotid artery for ~1 cm using micro-forceps, and clip the artery with an atraumatic microvascular clamp at the proximal end. At the distal end, ligate and cut off the artery.
9. Pass the distal end of the artery through a 26 G polyurethane barbed cuff (see Table of Materials) with a bevel end and grooves on the surface. Fix the artery with the cuff's handle using a microvascular clamp.
10. Remove the ligation suture at the distal end, turn the lumen inside out over the cuff, and fix with a superficial barb and grooves with a 10-0 surgical suture.
11. After preparing the recipient's vessels, drop 100 IU/mL of heparin saline on the vessels to prevent thrombosis. Cover the cervical incision with sterile wet saline gauze for subsequent implantation.

2. Donor procedure

1. Employ the same anaesthetic procedure (step 1.1) for the donor mouse.
2. Shave the abdominal hair using an electric razor, and disinfect the surgical area with three alternating rounds of povidone-iodine scrub followed by alcohol.
3. Incise the abdomen (2-3 cm) with a scissor along the midline from the symphysis pubis to the subxiphoid, and expand the incised area with a retractor.
4. Dissect 1 cm of the abdominal aorta and inferior vena cava using an electro-coagulator and a micro-forceps, and perform heparinization by injecting 1 mL of physiological saline supplemented with 250 IU/mL of heparin through the inferior vena cava. After this, excise the abdominal aorta and inferior vena cava.
5. Excise the thorax along the anterior axillary line on both sides using a surgical scissor to separate the chest wall. Ligate the superior vena cava with an 8-0 surgical suture.
6. Insert a scalp needle at the suprahepatic inferior vena cava. Then, inject ice-cold physiological saline supplemented with 100 IU/mL of heparin through the scalp needle from suprahepatic inferior vena cava to perfuse the donor heart until the blood color fades.
7. Re-perfuse the donor heart with 2-3 mL of ice-cold histidine-tryptophan-ketoglutarate (HTK) solution (see Table of Materials) using a scalp needle from the aortic arch to protect the donor myocardium. The mean warm ischaemia time is 5 min.
8. Ligate the superior and inferior vena cava and the pulmonary vein with a 5-0 surgical suture. Dissect and cut off the donor aorta and pulmonary artery before their branching. After that, divide the superior and inferior vena cava and the pulmonary vein to remove the donor's heart.

3. Implantation

1. Implant the donor heart into the cervical pocket of the recipient mouse in an inverted position.
2. Pull the cuff with an everted recipient jugular vein into the lumen of the donor pulmonary artery to perform end-to-end anastomosis of the donor pulmonary artery to the recipient external jugular vein. Ligate the cuff using the grooves on the surface through a 10-0 surgical suture to fix the anastomosis.
3. Employ a similar procedure for end-to-end anastomosis of the donor aorta to the recipient carotid artery.
4. Release the atraumatic microvascular clamp of the jugular vein followed by the carotid artery to re-perfuse the donor's heart. The mean cold ischaemia time is 15 min.
5. Fix the cardiac graft and suture it properly to prevent twisting of the graft.
6. Close the cervical incision with continuous sutures using a 5-0 polyamide monofilament suture (see Table of Materials).
   NOTE: Remove the suture after the wound is completely healed. 
7. Retain the recipient mouse inside a warm, dry, and clean cage until it recovers from anaesthesia.
   NOTE: It takes 5-10 min to recover.
8. Inject buprenorphine (0.05 mg/kg) subcutaneously into the recipient mouse every 6 h for 48 h for postoperative analgesia.
   NOTE: The analgesia dosage was optimized for this study. However, the analgesia regimen can be extended/ modified if there is any sign of pain in accordance with the institutional animal use guidelines. 

Representative Results

In this mouse cervical heterotopic heart transplantation model, the survival rate of recipient mice was approximately 95.2% (20 out of 21 mice survived). The primary cause of death was postoperative bleeding. The fast heartbeat with a regular rhythm serves as an indicator of the survival of the implanted donor heart.

C57BL/6 and BALB/c mice were MHC (H-2b) and MHC (H-2d) types in this model, respectively^7,8. These two strains differ by the H-2, which causes acute T-cell-mediated rejection⁹. Of all the cardiac allografts, 62.5% were lost within 7 days after transplantation, as assessed by palpating the heartbeat. All cardiac allografts were lost within 8 days after transplantation. In contrast, all the isogeneic heart transplants survived beyond 4 weeks (Figure 2). Mice that survived beyond 4 weeks were euthanized by CO₂ inhalation.

Figure 1: Schematic of the mouse cervical heterotopic heart transplantation model. (A) Protocol for preparing the recipient: after clipping the common carotid artery and external jugular vein at the proximal end, the vascular lumen of vessels is everted and fixed after passing through the barbed cuff with a bevel end and grooves on the surface. The dashed square shows the structure and usage of the cuff. (B) Donor heart resection: after the donor heart's perfusion with heparin and HTK solution from the inferior vena cava and aorta, the superior and inferior vena cava and pulmonary vein are ligated with sutures. The donor's heart is then resected by incising the vascular vessels. (C) Implantation of the donor's heart. The donor pulmonary artery and aorta is anastomosed to the recipient's external jugular vein, and carotid artery via the cuff with the recipient's vasculature turned inside out in an end-to-end pattern. Please click here to view a larger version of this figure.

Figure 2: Survival curve for cardiac grafts. The survival curve of cardiac grafts shows that allogeneic heart transplants are lost within 8 days after transplantation, which was assessed by palpation of the heartbeat. A total of 10 recipient mice underwent the modified cervical heterotopic heart transplantation in each group. All the isogeneic heart transplants survived more than 4 weeks. Please click here to view a larger version of this figure.

	Rupert Oberhuber et al.10	Xin Mao et al (present work)
Anaesthesia	xylazine and ketamine	isoflurane (Safe, continuous and stable)
Right lobe of the submandibular gland	removal	preservation (Reduce twirling of grafts)
Right sternocleidomastoid	removal	preservation (Reduce twirling of grafts)
Cardiac perfusion	Retrograde perfusion with 4 °C HTK solution from the aortic arch	1. Anterograde perfusion with ice-cold physiological saline supplemented with 100 IU/mL heparin solution from the suprahepatic vena cava. 2. Retrograde reperfusion with ice-cold HTK solution from the aortic arch. (Reduce coagulation and increase myocardial protection)
Cuff	blunt end, with handle	bevel end, with handle, barb and grooves on the surface (Facilitate eversion and fixation)

Table 1: Comparison of heart transplantation techniques. The current mouse cervical heterotopic heart transplantation technique is modified from Oberhuber, R. et al.¹⁰ and possesses additional advantages for cardiac graft survival.

Discussion

The mouse heart transplantation model contributes to the investigation of rejection mechanisms after heart transplantation, contributing to the development of unique approaches to improve the long-term survival of cardiac allograft recipients. However, heart transplantation in mice is a complex and challenging task, requiring a high level of microsurgery techniques, especially in vascular anastomosis^11,12,13. The mouse abdominal heterotopic heart transplantation model is performed using stitches by anastomosis of the donor aorta and pulmonary artery to the recipient aorta and inferior vena cava. The recipient's aorta and inferior vena cava need to be blocked in this operation. Therefore, ischaemia of the lower body and thrombosis of the inferior vena cava can increase the disability and death of recipient mice. To reduce the difficulties of vascular anastomosis during transplantation, Matsuura et al. first introduced a cervical heart transplantation model in mice using the cuff technique in 1991¹⁴. In this model, the oversleeve-like everted anastomosis of vessels by ligation with cuff increased the anastomosis efficiency. In contrast to the anastomosis of vessels with sutures in an abdominal heart transplantation mouse model, it reduced the bleeding probability post-procedure. Therefore, the improvement of the anastomosis efficiency reduced the ischaemia time of cardiac muscle and increased the survival rate of cardiac grafts. Additionally, cervical implantation of the donor's heart does not interrupt the circulation of the recipient aorta and inferior vena cava compared with abdominal implantation¹⁵; therefore, the survival of recipient mice is increased.

A unique experimental mouse heterotopic heart transplantation model is described here, established by Rupert Oberhuber et al.¹⁰. The procedure involves an end-to-end cervical anastomosis of the donor aorta and pulmonary artery to the recipient carotid artery and jugular vein, following a modified Cuff technique. In this model, the systemic circulation of the recipient mice does not interfere with¹⁰, and the donor's heart was perfused from the inferior vena cava and aorta with heparin and HTK solutions for better myocardial protection. However, the critical component of this model differed from that of Oberhuber et al.¹⁰, which employed the modified barbed cuff with a bevel end and grooves similar to that of Finsterer et al.¹⁶. The bevel end facilitates an oversleeve-like evert of the vascular lumen. The grooves on the surface facilitate the fixation of everted vessel walls with a cuff using sutures, and the barb outside the cuff reduces the slippage of anastomosed vessel walls from the cuff (Figure 1). These modifications shorten the surgical time by 20% and improve cardiac grafts' implantation efficiency and survival. Furthermore, the modified barbed cuff is produced from the most common polyurethane catheter used for scalp acupuncture, thus significantly reducing the cost of the procedure. A comparison of the current technique with that of Oberhuber et al.¹⁰ is shown in Table 1.

The unique features of this model need to be noted. First, the length and calibre of the cuff are essential for successful anastomosis. The suitable length of the cuff was ~3 mm with a handle (1 mm) (Figure 1). The suitable calibre of the cuff is 26 G and 22 G for artery and veins, respectively. The unsuitable length and calibre of the cuff would result in twisting or excessive tension of the anastomosed vessels. Second, the suitable lengths of the recipient's vessels are 1.5 to 2-folds of the cuff. Third, the donor's heart is not perfused with excessive pressure, potentially damaging the graft. Fourth, the cardiac graft is fixed, and the cuff is anastomosed in a suitable position by suturing after implantation to avoid movement or twisting of anastomosed vessels or grafts. Fifth, preservation of the submaxillary gland and sternocleidomastoid contributes to reducing twirling or twisting of the anastomosed vessels or graft when resecting the right sublingual gland to produce a fossa for cardiac graft. Sixth, to facilitate the oversleeve-like evert of the vascular lumenand reduce thrombosis after surgery, heparin solution (100 IU/mL) can be provided to the anastomosed vessels while performing anastomosis.

This cuff technique facilitates the anastomosis of donor and recipient vessels during implantation; however, the hardness of the cuff may, in turn, increase the risk of twisting the anastomosed vessels, resulting in an increase in thrombosis after transplantation. Optimization of the cuff material is warranted to reduce complications, increase graft survival, and increase the utilization rate of the models in subsequent experiments. Furthermore, the fibrous scar of the cut may limit the space for the cardiac graft and affect its long-term survival. In addition, ejection of the cardiac graft may interfere with recipient mice's normal haemodynamic blood flow. Finally, this model is non-functional and cannot be used to evaluate the cardiac function of grafts. Nevertheless, this study provides knowledge regarding heart transplantation's immunological and pathological functions.

Materials

Name	Company	Catalog Number	Comments
5-0 Polyamide Monofilament suture	B.Braun Medical Inc.	C3090954
8-0 Polyamide Monofilament suture	B.Braun Medical Inc.	C2090880
10-0 Polyamide Monofilament suture	B.Braun Medical Inc.	G0090781
22 G polyurethane cuff	B.Braun Medical Inc.	4251628-02
26 G polyurethane cuff	Suzhou Linhua Medical Instrument Co., LTD	REF383713
Anesthesia induction chamber	RWD Life Science Co., LTD	V100
Atraumatic microvascular clamp	Beyotime	FS500
BALB/c and C57BL/6 mice (20–30 g)	Centre of Experimental Animals (Army Medical University, Chongqing, China)
Buprenorphine	US Biological life Sciences	352004
Electrocoagulator	Guangzhou Runman Medical Instrument Co., LTD	ZJ1099
Gauze	Henan piaoan group Co., LTD	10210402
Heating pad	Guangzhou Dewei Biological Technology Co., LTD	DK0032
Heparin	North China Pharmaceutical Co., LTD	2101131-2
HTK solution	Shenzhen Changyi Pharmaceutical Co., LTD	YZB/Min8263-2013
Injection syringe (10 mL)	Shandong weigao group medical polymer Co., LTD	20211001
Isoflurane	RWD Life Science Co., LTD	21070201
Physiological saline	Southwest pharmaceutical Co., LTD	H50021610
Scalp needle	Hongyu Medical Group	20183150210
Shaver	Beyotime	FS600
Small animal anesthesia machine	RWD Life Science Co., LTD	R500
Surgical operation microscope	Tiannuoxiang Scientific Instrument Co. , Ltd, Beijing, China	SZX-6745
Swab	Yubei Medical Materials Co., LTD	21080274