Murine Cervical Aortic Transplantation Model using a Modified Non-Suture Cuff Technique

Summary

Here, we present a protocol of heterotopic aortic transplantation in mice using the non-suture cuff technique in a cervical murine model. This model can be used to study the underlying pathology of chronic allograft vasculopathy (CAV) and can help evaluate new therapeutic agents in order to prevent its formation.

Abstract

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. However, only minor improvement in the long-term results of transplanted solid organs could be observed over the past decades. In this context, chronic allograft vasculopathy (CAV) still represents the leading cause of late organ failure in cardiac, renal and pulmonary transplantation.

Thus far, the underlying pathogenesis of CAV development remains unclear, explaining why effective treatment strategies are presently missing and emphasizing a need for relevant experimental models in order to study the underlying pathophysiology leading to CAV formation. The following protocol describes a murine heterotopic cervical aortic transplantation model using a modified non-suture cuff technique. In this technique, a segment of the thoracic aorta is interpositioned in the right common carotid artery. With the use of the non-suture cuff technique, an easy to learn and reproducible model can be established, minimizing the possible heterogeneity of sutured vascular micro anastomoses.

Introduction

Over the past six decades, solid organ transplantation has evolved from an experimental procedure to a standard of care for the treatment of end-stage organ failure¹. Due to the improvement of antimicrobial agents, surgical techniques and advancement in immunosuppressive regiments, the early success rate of solid organ transplantation have significantly increased over the past decades².

However, long-term graft survival rates have not significantly improved in the same manner³. The development of CAV is the major factor limiting long-term survival^4,5,6. This pathology is characterized by the formation of a concentric neointimal layer consisting of smooth muscle cells, leading to progressive narrowing of the vessel and consecutive malperfusion of the transplanted solid organ. In heart transplant recipients, CAV lesions can be diagnosed in up to 75% of patients 3 years after transplantation⁷.

The pathophysiology of CAV is not fully understood yet. It seems to be related to numerous immunological and non-immunological factors, leading to endothelial damage with subsequent endothelial activation and dysfunction⁸. Thus far, no causal treatment option exists for the prevention of CAV, emphasizing the need for a reproducible small animal model in order to study the formation and potential therapy of CAV.

With the use of murine aortic transplantation models, CAV like lesions can be seen 4 weeks after transplantation. Those lesions consist mainly of vascular smooth muscle cells, thereby, resembling the human pathology. Because of a wide variety of transgenic and knock out mice, the use of mouse models in transplant associated pathologies offers a unique opportunity to identify new therapeutic options and understand their development. Due to the small diameter of the transplanted vessels however, the use of mouse models is commonly associated with long learning curves and an initial high complication rate⁹. With the introduction of the non-suture cuff technique, this most challenging part of the operation can be facilitated and the diameter of the anastomosis is kept constant^10,11.

Protocol

All experiments were performed according to the guidelines of the German animal welfare act (TierSchG.) (AZ: 55.2-1-54-2532.Vet_02-80-2015).

1. Animal housing

1. For experiments, use male C57BL/6 and BALB/c mice weighing 20-25 g with C57BL/6 mice as the recipient animals and BALB/c mice as the donor animals.
2. Purchase the animals and house in a barrier pathogen-free facility, in accordance with the FELASA guidelines for health monitoring¹².
3. Keep the mice in standard Makrolon cages with enrichment nesting material. Provide ad libitum access to water and pelleted food at a day/night cycle of 12 h.
4. Maintain the room temperature at 22 ± 2°C and the relative humidity at 55 ± 5%.

2. Recipient preparation

1. First, anesthetize the recipient animal (C57BL/6) with an intraperitoneal injection of midazolam (5 mg/kg; 5 mg/mL), medetomidin (0.5 mg/kg; 1 mg/mL) and fentanyl (0.05 mg/kg; 0.05 mg/mL).
   NOTE: The correct depth of the anesthesia should be reached in 5-10 min.
   1. Pinch the hind feet with forceps to check for reflexes to confirm the depth of anesthesia.
2. Clip all the hair of the cervical lateral region with an electric hair clipper for small animals and apply ophthalmic ointment with cotton swabs to prevent the eyes from drying out during the procedure.
3. Place the animal in a supine position on a heating pad under the microscope and gently tape its legs to the operating table with skin sensitive plaster strips.
4. Tilt its head back and scrub the operative field several times with alcohol.
5. Make a skin incision from the jugular incision to the right lower mandible with small scissors.
6. Remove the right lower lobe of the submandibular gland via bipolar cautery of the vessel pedicle and subsequent excision with microscissors.
7. Remove the right sternocleidomastoid muscle via bipolar cautery of the upper and lower portion and subsequent excision with microscissors to gain access to the common carotid artery.
8. Mobilize the common carotid artery as far distally and proximally as possible by pulling the surrounding connective tissue apart with fine forceps.
9. Tie two 7-0 silk ligatures with minimal distance between each around the middle of the common carotid artery and transect the common carotid artery with fine microscissors between the ligatures.
10. Pass the proximal, ligated end through the cuff and fix it with a small artery clamp.
    NOTE: The cuffs that were used in this procedure were cut with microscissors from tubes of polyimide with an outer diameter of 0.610 mm and a wall thickness of 0.0254 mm. The completed cuffs had a length of ~2 mm with one half, which is used for the cuffing process, being a full cylinder and the other half, which is clamped, being a half-cylinder.
11. Remove the ligature with fine microscissors, as close to the ligature as possible, and flush the lumen with heparinized saline (50 IU/mL) with a 30 G needle, while taking care not to damage the vessel walls.
12. Distend the open lumen using fine vascular dilatators and evert the carotid stump over the cuff by pulling it gently over the polyimide tube.
13. Immediately fix the everted carotid with a loosely pre-tied 7-0 silk loop.
    NOTE: Loosely pre-tie 4 7-0 silk loops with a diameter of about 1.5 mm before the surgery to make the cuffing-procedure smoother and easier.
14. Perform the same procedure (2.10-2.13) at the other end of the carotid artery.
15. Set the recipient animal aside and moisturize the operative field with saline until the aortic segment is explanted.

3. Donor operation

1. Anesthetize the donor mouse (BALB/c) in the same fashion as the recipient animal.
   1. Pinch the hind feet with forceps to check for reflexes to confirm sufficient anesthesia.
2. Clip all hair of the abdominal and thoracic region with an electric hair clipper for small animals and apply ophthalmic ointment with cotton swabs to prevent the eyes from drying out during the procedure.
3. Place the animal in a supine position on a heating pad under the microscope and gently tape its legs to the operating table with skin sensitive plaster strips.
4. Scrub the operative field several times with alcohol.
5. Perform a midline abdominal laparotomy with small scissors and push the intestines slightly upward to expose the inferior vena cava (IVC).
6. Inject the inferior vena cava (IVC) with 1 mL of heparinized saline using a 30 G needle.
7. Cut the abdominal aorta and IVC below the renal arteries with small scissors to exsanguinate the donor animal. Loosely place a compress into the abdomen to absorb the blood.
8. Perform a thoracotomy at the bilateral diversion of the ribs with scissors and tilt the anterior chest wall cranially with a surgical clamp to expose the mediastinum.
9. Cut the IVC and the esophagus directly above the diaphragm with microscissors.
10. Remove the heart and the lungs by tilting them upward with forceps holding the cut IVC/esophagus and then excising them with microscissors from their base to gain access to the thoracic aorta in the dorsal mediastinum.
11. Mobilize the thoracic aorta from its surrounding tissue by pulling apart the surrounding connective tissue and fat with fine forceps while being careful not to damage any intercostal arteries.
12. Cauterize all branches from the thoracic aorta with bipolar cautery forceps and excise the aortic segment between the diaphragm and the aortic arch using microscissors.
13. Flush the excised aortic segment with heparinized saline with a 30 G needle, while taking care not to damage the vessel walls, to remove any remaining blood or clots, and transfer the graft to the recipient animal.
    NOTE: Directly place the aortic graft in the roughly right position in the recipient during transfer. If there are problems confusing the different ends of the graft in the recipient animal, a loose ligature around the distal end could help.

4. Implantation

1. Pull the proximal end of the donor aortic segment over the proximal cuff on top of the everted carotid artery with fine forceps and immediately fix it with a loosely pre-tied 7-0 silk loop.
2. Trim the distal, free end of the aortic graft with microscissors so that the graft length fits the distance between the two cuffs.
3. Repeat step 4.1 on the other end of the aorta with the other cuff to complete the anastomosis.
4. Remove the distal clamp to allow retrograde perfusion.
5. After achieving hemostasis, remove the proximal clamp to complete the anastomosis.
6. Finally, close the wound with 6-0 continuous suture.

5. Postoperative care

1. Monitor the mouse closely in the first 6 h after the operation and then several times a day for the first 72 h after the transplantation to detect any complications instantly.
2. For postoperative analgesia, inject the transplanted mouse with buprenorphine (0.05-0.1 mg/kg) subcutaneously directly after the transplantation and then every 12 h for 72 h to provide appropriate, long term analgesia.

6. Aortic graft explanations

1. Anesthetize the transplanted animal with an intraperitoneal injection of midazolam (5 mg/kg; 5 mg/mL), medetomidin (0.5 mg/kg; 1 mg/mL) and fentanyl (0.05 mg/kg; 0.05 mg/mL) 4 weeks after transplantation.
   1. Pinch the hind feet with forceps to check for reflexes to confirm sufficient anesthesia.
2. Clip all hair of the abdominal, thoracic and cervical region with an electric hair clipper for small animals.
3. Place the animal in a supine position on a heating pad under the microscope and gently tape its legs to the operating table with skin sensitive plaster strips.
4. Scrub the operative field several times with alcohol.
5. Perform a midline abdominal laparotomy with small scissors and push the intestines slightly upward to expose the inferior vena cava (IVC).
6. Inject the inferior vena cava (IVC) with 1 mL of heparinized saline using a 30 G needle.
7. Cut the abdominal aorta and IVC below the renal arteries with small scissors to exsanguinate the donor animal. Loosely place a compress into the abdomen to absorb the blood.
8. Make a skin incision from the jugular incision to the right lower mandible with small scissors corresponding to the skin incision made during the transplant procedure.
9. Identify the transplanted aortic graft together with the distal and proximal cuff and blunt remove any surrounding tissue with forceps.
10. Using microscissors, cut through the common carotid artery distal and proximal to the aortic graft with the cuffs in order to explant the aortic graft together with the two cuff ends.
11. Cut the aortic segment in half and preserve the specimens for further analyses (frozen sections, paraffin embedded sections, snap frozen material)^13,14.

Results

In the fully MHC-mismatch transplantation model, a concentric neointimal layer can be seen 4 weeks after transplantation (Figure 2). This layer consists primarily of vascular smooth muscle cells as immunohistological staining for SM22 (a selective marker for mature vascular smooth muscle cells) revealed. As stated before, these vascular smooth muscle cells are pathognomonic for lesions seen in chronic allograft vasculopathy. For further analyses, aortic segments should be sectioned and stain...

Discussion

Chronic allograft vasculopathy is the major cause of late graft loss after solid organ transplantation of the heart and likely renal and lung allografts⁸. Thus far, no causal therapeutic regimen could be developed in order to prevent the formation of CAV.

The pathophysiology of CAV is multifactorial and involves immunological and non-immunological aspects¹⁶. The use of rodent models in transplantation have been essential in understanding the unde...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Balb-c Mice (H2-d)	Charles River	Strain# 028	Donor animal
Bipolar cautery system	ERBE	ICC 50 / 20195-023	Bipolar cautery
C57BL/6J (H-2b)	Charles River	Strain# 027	Recipient animal
Halsey Needle Holders	FST	12501-12	Needle Holder
Halsted-Mosquito Forceps	AESCULAP	BH111R	Curved Clamp
Medical Polyimide Tubing	Nordson MEDICAL	141-0031	Cuff-Material
Micro Serrefines	FST	18055-04	Micro Vessel Clip
Micro-Adson Forceps (serrated)	FST	11018-12	Standard Forceps
Micro-Serrefine Clamp Applying Forceps	FST	18057-14	Clipapplicator
S&T Forceps - SuperGrip Tips (Angled 45°)	S&T	00649-11	Fine Forceps
S&T Vessel Dilating Forceps - Angled 10° (Tip diameter 0.2 mm)	S&T	00125-11	Vesseldilatator
Schott VisiLED Set	Schott	MC 1500 / S80-55	Light
Stereoscopic microscope	ZEISS	SteREO Discovery.V8	Microscope
Student Fine Scissors / Surgical Scissors - Sharp-Blunt	FST	91460-11 / 14001-12	Standard Sissors
Vannas-Tübingen Spring Scissors (curved, 8.5 cm)	FST	15004-08	Microsissors (curved)
Vannas-Tübingen Spring Scissors (straight, 8.5 cm)	FST	15003-08	Microsissors (straight)