Modified Heterotopic Hindlimb Osteomyocutaneous Flap Model in the Rat for Translational Vascularized Composite Allotransplantation Research

Summary

Vascularized composite allograft offers life-altering benefits to transplant recipients, but the biological causes of graft rejection and vasculopathy remain poorly understood. The rodent surgical model presented here offers a reproducible, clinically relevant model of transplantation, allowing researchers to evaluate rejection events and potential therapeutic strategies to prevent their occurrence.

Abstract

Vascularized composite allotransplantation (VCA) is a relatively new field in the reconstructive surgery. Clinical achievements in human VCA include hand and face transplants and, more recently, abdominal wall, uterus, and urogenital transplants. Functional outcomes have exceeded initial expectations, and most recipients enjoy an improved quality of life. However, as clinical experience accumulates, chronic rejection and complications from the immunosuppression must be addressed. In many cases where grafts have failed, the causative pathology has been ischemic vasculopathy. The biological mechanisms of the acute and chronic rejection associated with VCA, especially ischemic vasculopathy, are important areas of research. However, due to the very small number of VCA patients, the evaluation of proposed mechanisms is better addressed in an experimental model. Multiple groups have used animal models to address some of the relevant unsolved questions in VCA rejection and vasculopathy. Several model designs involving a variety of species are described in the literature. Here we present a reproducible model of VCA heterotopic hindlimb osteomyocutaneous flap in the rat that can be utilized for translational VCA research. This model allows for the serial evaluation of the graft, including biopsies and different imaging modalities, while maintaining a low level of morbidity.

Introduction

Reconstructive surgery for the catastrophic tissue loss from amputation, blast injuries, malignancies, and congenital defects are limited by the availability of tissue from the patient and the additional morbidity caused at the donor site. In some cases, such as burn victims or quadrilateral amputees, viable tissue for reconstruction is not available from the patient. In 1964, the first modern hand transplant was performed in Ecuador. While this was a technical success, immunosuppression available at the time was insufficient to prevent rejection, and the graft was lost in less than 3 weeks¹. In 1998 and 1999, the first hand transplants in the modern era of immunosuppression were performed in Lyon, France² and Louisville, Kentucky, USA³. For the first time, reconstructive surgeons could replace like with like. Face transplantation was first performed in 2005⁴, and a number of other VCA grafts are now routinely being performed, such as abdominal wall⁵, uterine, and urogenital transplants⁶.

Unlike solid organ transplantation, most VCA techniques involve the presence of the highly antigenic donor skin. Clinical experience has determined that the acute rejection of the skin is relatively easy to control but may contribute to the chronic rejection of the underlying tissues and vessels, which do not respond well to treatment⁷. The vascular dysfunction associated with an alloimmune response is a more ominous obstacle for the field of VCA⁷. Macrovasculopathies lead to perfusion deficits, delayed healing, and proinflammatory conditions. Both confluent aggressive large-vessel vasculopathy and focal intimal hyperplasia occur in hand transplant recipients⁷. Additionally, microvasculopathies likely contribute to VCA complications as well and may even lead to rejection events. While both immune and nonimmune factors likely play a role in the vasculopathy of hand transplant recipients, the specific mechanisms promoting distal vessel dysfunction in VCA are not known, particularly in the context of low-grade, chronic rejection. These unanswered questions necessitate the development of an animal VCA model that will allow for the serial assessment of the graft during the clinical course of VCA rejection/maintenance and vasculopathy. Such a model will offer insights into the rejection and vasculopathy in the face of immunosuppression, infectious challenge, and/or other postoperative traumatic injury^8,9.

Presented here is an allogeneic rat VCA heterotopic hindlimb osteomyocutaneous flap model. Based on previously published VCA models, this procedure is technically easy to perform, reproducible in a large number, and exhibits minimal morbidity and discomfort to the recipient animal. This model was designed to allow clinical and histopathological assessments of VCA acceptance vs. rejection, and provides an opportunity to evaluate underlying immune and nonimmune mechanisms involved in rejection.

Protocol

All animal surgeries were performed in accordance with protocols approved by the University of Louisville's Institutional Animal Care and Use Committee (IACUC-approved protocol 18198) and the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals¹⁰. Four-month-old male Brown-Norway (RT1.Aⁿ) and 4-month-old male Lewis (RT1.A^l) rats were used as VCA donor and recipients, respectively.

1. Donor Allograft Harvest

1. Sedate the donor animal using vaporized isoflurane applied through a chamber.
2. Shave the graft donor area (hindlimb), as well as the groin and abdomen areas. Following that, treat with depilatory cream in order to reduce the amount of fuzz left by the clippers.
3. Deeply anesthetize donor animals using intraperitoneal (IP) ketamine (60 mg/kg)/xylazine (15 mg/kg)/acepromazine (2 mg/kg). Administer an initial dose of 0.2 mL/100 g of body weight and additional doses of 0.2 mL every h. For convenience, it is optional to perform this step before step 2.
4. Continuously monitor animals while under anesthesia for respiration, body temperature, and depth of anesthesia, using the toe pinch withdrawal reflex test.
5. Administer 30 U of heparin solution subcutaneously (SC) in the scruff area prior to the surgery to prevent clotting.
6. Wear a mask, a head cover, a disposable isolation gown, and disposable gloves.
7. Place the donor animal supine on a heating pad. Produce a sterile surgical field by prepping, scrubbing, and draping the surgical area including the both ventral and dorsal aspects of the leg. Don sterile gloves.
8. Make a 3 cm skin incision in the groin concavity using scalpel blade #15 and reflect the inguinal fat pad laterally using iris scissors.
9. Expose the common femoral vessels and place a wire hook with an elastic band to retract the abdominal muscles.
10. Using a dissecting microscope (40x), dissect the pedicle proximally from the emergence of the common femoral vessels under the inguinal ligament and distally to the confluence of popliteal vessels into the graft.
11. Using microclips and bipolar jewelers’ forceps, ligate and divide the large arterial and venous branches, such as lateral circumflex femoral vessels, superficial caudal epigastric vessels, the saphenous artery, and proximal caudal femoral vessels, to mobilize the main femoral vessels. Cauterize any small branches using fine bipolar forceps.
12. Make a skin incision from the center of the previous skin cut along the ventral side of the hindlimb, to the ankle area, using iris scissors.
13. Cut the gracilis muscle, as well as the other adductor muscles underneath it, in a vertical fashion to expose and ligate the medial proximal genicular vessels, deep-branching small vessels, and the sciatic nerve.
    NOTE: At this point, on a separate surgical table, the other surgeon should intubate and anesthetize (2.5%–3% isoflurane) the recipient animal; this allows the surgeons to prepare the recipient surgical site in time for graft placement and minimize the graft's ischemic time.
14. On the donor animal, make circumferential skin incisions at the level of the knee and ankle. Disarticulate the knee and ankle, remove extraneous muscle and tissue, and make a vertical skin incision on the dorsal side of the hindlimb to free the graft. At this point, the graft (composed of fibula and tibia, covered with related muscles and skin island nourished by its perforators) is connected only by the pedicle.
15. Place small clamps as proximally as possible on the femoral artery and vein, and cut the pedicle as proximally as possible, near the inguinal ligament.
16. To flush the graft of blood, inject heparinized saline (30 U/mL) into the femoral artery using a 27 G flushing blunt cannula.
    NOTE: Dilating the artery prior to the heparin flush allows easy access for insertion of the cannula. During the flush, closely monitor the outflow from the femoral vein. Once clear fluid exits the femoral vein, stop the flush.
17. Wrap the isolated graft in warm saline-soaked gauze and transport it immediately to the recipient animal’s table. At this time, the recipient surgical site should already be prepared for vascular anastomosis.
18. After the graft harvest, immediately euthanize the donor rat via pneumothorax.

2. Recipient Transplantation Surgery

1. Following the sedation induction using vaporized isoflurane applied through a chamber, deeply anesthetize the recipient animal via a ventilator-controlled endotracheal tube and 2.5%–3% isoflurane.
   NOTE: At this stage, the donor rat is still anesthetized.
2. Continuously monitor the heart rate, respiratory rate, body temperature, and depth of anesthesia of the recipient animal, using the toe pinch withdrawal reflex test.
3. In order to prevent dehydration and hypoglycemia, inject 2 mL of lactated Ringer’s solution and 2.5% dextrose subcutaneously at the beginning and another 2 mL at the end of the surgery.
4. Shave the groin area, then treat with depilatory cream in order to reduce the amount of fuzz left by the clippers.
5. Wear a mask, a head cover, a disposable isolation gown, and sterile gloves.
6. Place the animal supine on a heating pad. Apply ophthalmic ointment to prevent corneal abrasions during anesthesia. Produce a sterile surgical field by prepping, scrubbing, and draping the surgical area.
7. Make a 3 cm skin incision in the groin concavity using scalpel blade #15 and reflect the inguinal fat pad laterally using iris scissors.
8. Expose the common femoral vessels and place a wire hook with an elastic band to retract the abdominal muscles.
9. Ligate and divide Murphy branches.
10. Using 10-0 nylon interrupted sutures, anastomose donor vessels to recipient vessels via venous end-to-side technique and arterial end-to-end technique. Gradually release the clamps from the artery and then the vein. Monitor the anastomotic sites for bleeding and add additional sutures if needed.
11. Visually assess the vascular anastomosis in order to assure effective graft reperfusion.
12. Inset the graft into the inguinal pocket and orient it upside down, with the ankle joint superior and knee joint inferior.
13. Using tucking sutures, secure the graft to adjacent muscles. Close the skin via interrupted horizontal mattress skin absorbable 4-0 sutures.
14. Remove the recipient animal from anesthesia and wean it off the ventilator. Place the animal on a heating pad for thermal support.
    NOTE: The overall operation time is between 3 to 4 h, depending on the surgeon’s experience and acquaintance with the surgical procedure.
15. Administer meloxicam (1 mg/kg) subcutaneously for pain suppression and monitor until the animal is fully recovered and mobile.

3. VCA recipient Monitoring

1. House the recipient rats singly and monitor them daily for clinical signs of pain, dehydration, weight loss, and decreased activity in addition to surgical failure (for the first 48–72 h) or rejection. Administer meloxicam subcutaneously (1 mg/kg) daily for the first 3 days for pain suppression.
2. Based on the research endpoint, choose an immunosuppressant drug to be administered.

4. Histology

1. Under inhaled isoflurane anesthesia (2.5%–3%), obtain the serial skin and underlying muscle biopsies from the donor graft at desired time points. The skin should be scrubbed and draped prior to obtaining a biopsy, and a sterile field and technique should performed.
2. Close the wound with one to two stitches, using absorbable 4-0 sutures. Return the animal to its cage and allow it to recover from the anesthesia.
3. Fix the biopsied tissues in separate tubes in 10% formalin.
4. At the terminal time point and under inhaled isoflurane anesthesia (2.5%–3%), take a larger skin biopsy that spans the donor/recipient border. Carefully locate the vessel leash pair at the site of anastomoses; the proper site will be apparent due to the sutures. Take the desired vessel samples from the artery and/or vein. Fix all samples separately in 10% formalin. After collection of tissue samples, and while the animal is still under isoflurane anesthesia, immediately euthanize the animal via pneumothorax.
5. Using a tissue processor (or other preferred embedding technique), paraffin-embed each biopsy into its own block. For skin samples, orient the tissue so that all epidermal and dermal layers may be seen in a single slice. For vessel samples, orient the vessels so that cross sections may be obtained.
6. Using a microtome, cut 6 µm-thick sections and apply them to slides for hematoxylin and eosin (H&E) staining.
7. Stain for H&E using a standard protocol.
8. Obtain representative images of all desired tissue samples using brightfield microscopy techniques.

Results

The rat VCA heterotopic hindlimb osteomyocutaneous flap model allows for long-term allograft survival under immunosuppression. The model is reliable, reproducible, and simple to perform. The flap is well hidden in the groin area and constitutes minimal morbidity and discomfort to the animal. The skin presentation is a clinical manifestation of the allograft’s survival and rejection (Figure 1). The flap design allows for gross clinical monitoring and creates an opportunity for various i...

Discussion

In developing this model of VCA, several key issues were considered. First, it was important to include intact bone (tibia and fibula), bone marrow, and skin in the graft. While clinical hand transplants from adult donors do not transfer significant amounts of actively hematopoietic marrow, studies of the role of the bone marrow niche are better mirrored using an intact, vascularized bone rather than a cut long bone, which results in fibrosis of the exposed marrow. Moreover, the closed bone osteomyocutaneous flap design ...

Materials

Name	Company	Catalog Number	Comments
Acepromazine	Henry Schein	5700850
Adventitia Scissors	ASSI	SAS15R8
Approximator Clamp (Double)	ASSI	ABB2V, ABB22V
Approximator Clamp (single)	FST	00398-02
Clamp Applying Forceps	ASSI	CAF4
Dissecting Scissors	ASSI	SDS18R8
Flushing blunt needle 27 G	SAI
Heparin Sodium	Sagent	25021-400-30
Isoflurane	Patterson Veterinary	14043-704-06
Jewelers Bipolar	ASSI	103000BPS03
Jewelers forceps #3	FST	11231-30
Ketamine HCl 100 mg/mL	Zoetis	043-304	DEA License required
Lactated Ringer Solution	Hospira	0409-7953-03
Lactated Ringer Solution + 5% Dextrose	Hospira	0409-7953-09
Meloxicam	Henry Schein	11695-6925-2
Micro forceps	ASSI	JFAL3
Micro needle holder	ASSI	B138
Prograf (Tacrolimus) 5 mg/mL	Astellas	0469-3016-01
Suture, 10-0 Prolene	Ethicon	W2790	or 10-0 Ethilon (2830)
Suture, 4-0 Coated Vicryl	Ethicon	J714D
Vessel Dilator Forceps	ASSI	D5AZ
Xylazine	VetOne	13985-612-50