Orthotopic Rat Forelimb Transplantation

Summary

Here, we present a protocol to demonstrate the microsurgical feasibility of rat forelimb transplantation. This model may serve as an important translational platform for VCA research.

Abstract

Vascularized composite allotransplantation (VCA) involves the transplantation of multiple tissue types -- including skin, muscle, bone, and nerves -- offering a promising reconstructive option for patients with severe traumatic injuries or disfigurements. Despite its transformative potential, VCA has encountered significant challenges such as graft rejection, chronic immunosuppression complications, and neuromuscular recovery's intricacies.

We utilize a rat forelimb model as a cost-effective and anatomically relevant platform to address these challenges. The rat forelimb closely mirrors human limb anatomy, enhancing our findings' translational impact. Previous studies have validated this model for reliably and reproducibly measuring functional recovery, thereby establishing it as a key tool for assessing the rejection trajectory of forelimb grafts.

Moreover, the model offers a valuable opportunity to explore innovative therapeutic approaches and serve as a good translational platform for novel preservation techniques. Through further investigation of this model, we aim to deepen our understanding of the mechanisms behind graft rejection and neuromuscular recovery. Ultimately, this work strives to pave the way for improving clinical outcomes of VCA, addressing both current limitations and future challenges in transplant medicine.

Introduction

Vascularized Composite Allotransplantation (VCA) is a life-changing procedure to reconstruct severe tissue defects. VCA refers to the transplantation of a composite tissue consisting of skin, muscle, bones, connective tissue, blood vessels, and nerves. Further refinement and exploration of VCA techniques would profoundly benefit patients suffering traumatic injuries or disfigurements¹. However, its broader application is challenged by ongoing hurdles such as graft rejection, complications from long-term immunosuppression, neuromuscular recovery, and peri-transplant logistics.

The rat forelimb model has emerged as a preferred choice for VCA studies due to its cost-effectiveness and its close resemblance to human anatomical and neuromuscular characteristics^2,3. Our previous work has demonstrated that this model provides a reliable and reproducible means of assessing behavioral functional recovery⁴. In this study, we present a detailed, step-by-step guide to the essential microsurgical techniques required for this model.

As VCA research continues to evolve, the rat forelimb model will allow mechanistic investigation of the rejection process, nerve regeneration, and neuromuscular functional recovery. Researchers could explore new preservation techniques, evaluate the effectiveness of machine perfusion methods, and test emerging immunomodulatory therapies, which may help reduce current reliance on long-term immunosuppression.

Protocol

Surgical procedures were carried out under the protocols approved by the Johns Hopkins University Animal Care and Use Committee (RA18M74) according to guidelines established by the National Institutes of Health and the American Association for the Accreditation of Laboratory Animal Care.

1. Donor procedure (Figure 1)

1. Conduct the induction of anesthesia via inhalation of isoflurane at a concentration of 5% and maintain at 1% throughout the procedure. Start the procedure after confirming the accurate anesthesia level via toe pinch.
2. Make a circumferential incision about 1 cm above the elbow. Divide the skin and expose the underlying superficial subcutaneous tissue without damaging it (Figure 1A).
3. Dissect the subcutaneous tissue/fascia, cauterize all superficial vessels, and transect. Subsequently, the deltoid, part of the pectoralis major, and the triceps brachii will be exposed.
4. Bluntly dissect beneath the pectoralis major muscle, revealing the brachial vessels, as well as the median and ulnar nerve (Figure 1B).
5. Cauterize the superficial vein on the deltoid muscle and transect proximally, then transect the deltoid muscle perpendicular to the muscle fiber orientation. Afterward, dissect and transect the pectoralis major and minor, revealing the axillary artery, vein, and nerve. (Figure 1C).
6. Transect the median, ulnar, and radial nerve at their proximal end (right after the split), preserving as much length as possible.
7. Dissect and free the brachial artery and vein.
8. Transect the biceps brachii, triceps brachii, and the latissimus dorsi muscle. Ensure that the humerus is properly exposed upon completion of dissection.
9. Use a rongeur forceps to transect the bone distal to the deltoid tuberosity of the humerus.
10. Gently check the lumen of the brachial artery and vein (Figure 1C). Flush the graft with 6 mL heparinized saline.
11. Mount polyimide cuff on both brachial artery and vein, respectively (Figure 1D).
12. Wrap the graft with wet cotton gauze and store it at 4 °C.
13. Euthanize the donor rat via an inhalant anesthetic (isoflurane) overdose. Once completely unresponsive to a toe pinch, perform a thoracotomy and excise the heart.

2. Recipient procedure (Figure 2)

1. Conduct the induction of anesthesia via inhalation of isoflurane at a concentration of 5% and maintain at 1% throughout the procedure. Confirm the status of the anesthesia via toe pinch.
2. Make a circumferential incision about 1-2 cm above the elbow.
3. Dissect the subcutaneous vessels and cauterize larger vessels to expose the deltoid, part of the pectoralis major, and the triceps brachii muscle.
4. Perform a blunt dissection beneath the pectoralis major muscle to reveal the brachial vessels, as well as the median and ulnar nerve.
5. Transect the deltoid and part of the pectoralis major and minor muscle at their distal end, exposing the brachial vessels, median, and ulnar nerves (Figure 2A).
6. Dissect and transect the median and ulnar nerve at their distal end. Ligate all vascular branches (Figure 2B).
7. Transect the biceps brachii, triceps brachii, and latissimus dorsi muscle to expose the humerus. Carefully cauterize any bleeding from the transected muscles.
8. Transect the humerus at the deltoid tuberosity. Use bone wax to seal the marrow cavity.
9. Use an intramedullary rod to reconnect the bone marrow cavity of the donor and recipient, then use bone wax to seal the connection. Suture the triceps brachii muscle with a 6-0 absorbable suture, thus creating a platform for vessel anastomosis.
10. Perform epiperineurial neurorrhaphy with interrupted 10-0 suture of the radial nerve.
11. Perform the vascular anastomosis using a non-suture cuff technique (Figure 2C).
12. Perform epiperineurial neurorrhaphy with interrupted 10-0 sutures to reconnect the median and ulnar nerves (Figure 2D).
13. Reconnect the deltoid and pectoralis major muscle using interrupted sutures.
14. Close the wound with 4-0 non-absorbable suture.
15. Administer warm, sterile saline at 2% of the body weight subcutaneously at the end of surgery.
16. Observe the animal for at least 4 h under a warming lamp for recovery. Return it to the animal housing facility once it is alert, can upright itself, demonstrate coordinated movement, and independently obtain water and food.

3. Long term post-transplant management

1. Monitor animals every day for signs of pain and distress as defined below.
   1. Check for anorexia indicated by the absence of feces in the cage.
   2. Check if the animal does not drink water, leading to dehydration evidenced by the tenting of the skin.
   3. Check if the animal is hunched up, unwilling to move, favoring a limb, or guarding the incision site.
   4. Check if the animal fails to groom, reflecting in a ruffled or dirty coat.
   5. Check if the animal excessively licks/scratches. Check for redness and swelling at the incision site and self-mutilation.
   6. Check for aggressive behavior, especially when attempting to pick up the animal.
   7. Check if the animal is panting, has labored breathing, and has reddish-brown nasal/ocular discharge.
   8. Check for sepsis.
2. Remove the sutures at 14 days post-operation.
3. Administer a second dose of buprenorphineSR (rat dose: 1.0 mg/kg SC) on postoperative day 2 (POD2; 48 h after the initial dose).
4. Start a weekly functional evaluation in week 4 post-transplantation.
   1. Briefly, under general anesthesia, fix the animal's forelimb in 90° abduction. Place two needle electrodes 1 mm apart in the axilla near the insertion of the pectoralis major onto the humerus.
   2. Stimulate the proximal median nerve percutaneously at 10 V to induce a maximal tetanic contraction of the flexors.
   3. Once full digital flexion forms a grasp, place a metal loop connected to a force meter on the palm and secure it within the flexed digits. Then, pull the force meter until the grasp is lost, recording the maximum force (in Newtons).

Results

Graft survival can be monitored and determined through macroscopic observation. Representative pictures of both short-term (POD7) and long-term follow-up (POD120) are shown in Figure 3. The graft could recover without apparent signs of necrosis. Survival data are shown in Figure 4. Both skin and muscle samples are harvested at the designated endpoint. Representative histological images are shown in Figure 5 and Figure 6. Both the figures exhibit a normal histology feature of skin and muscle. The grip strength test was applied to evaluate the functional recovery (mainly media nerve, Figure 7 and Figure 8). The recovery would reach a plateau around 15 weeks post-transplant. The technique has been extensively described in a previous publication⁵.

Figure 1: Donor procedure. (A) Circumferential incision; (B) exposing brachial vessels, median and ulnar nerve; (C) lumen check before flush; (D) non-suture cuff technique. (*brachial artery, @brachial vein, #median nerve, &ulnar nerve). Please click here to view a larger version of this figure.

Figure 2: Recipient procedure. (A) Exposing brachial vessels, median, and ulnar nerve; (B) dissection of brachial vessels, median, and ulnar nerve; (C) non-suture cuff technique for vessel anastomosis; (D) Epiperineurial neurorrhaphy. (*brachial artery, @brachial vein, #median nerve, &ulnar nerve). Please click here to view a larger version of this figure.

Figure 3: Forelimb graft. Representative macroscopic images of forelimb graft on post-transplantation day 7 (left) and day 120 (right). Please click here to view a larger version of this figure.

Figure 4: Graft survival. Please click here to view a larger version of this figure.

Figure 5: Hematoxylin and eosin staining of graft skin. The section shows normal histology features of the skin. Please click here to view a larger version of this figure.

Figure 6: Hematoxylin and eosin staining of graft muscle. The section shows normal histology features of the muscle. Please click here to view a larger version of this figure.

Figure 7: Demonstration of grip strength test. Please click here to view a larger version of this figure.

Figure 8: Recovery of grip strength post-transplantation. Please click here to view a larger version of this figure.

Discussion

Orthotopic forelimb transplantation in rats is a commonly used model in experimental reconstructive transplantation as it not only contains vascularized composite tissues but also allows for the assessment of functional recovery. A conventional suture technique for vascular anastomosis is highly complex and demands advanced microsurgical skills that require years of microsurgical training. To greatly reduce the time and costs, a vascular non-suture cuff anastomosis technique was applied in this modified rat forelimb model. It significantly expedites the microsurgical training process and shortens the training curve, as we have demonstrated in the mouse heart transplantation model⁶ and mouse hindlimb model⁷. The non-suture cuff technique has achieved over 90% success in all the forelimb transplant-related projects. From a technical perspective, the inner membranes of the vessel walls are free from surgical manipulation, which would help reduce the confounding factor in static cold storage-related experiments.

With the establishment of this model⁴, we have furthermore effectively tailored grip strength tests and the Irvine, Beatties, and Bresnahan scale (IBB scale) to measure functional recovery in the postoperative course. In addition, the model facilitates consistent histological assessments of peripheral nerve regeneration and muscle atrophy and is therefore ideally suited for a wide range of innovative studies like novel preservation techniques such as machine perfusion⁸, among others^9,10. The availability of a robust, highly reproducible model sets the stage for significant advancements in the field of VCA, benefiting both research endeavors and future clinical applications.

We would also like to acknowledge the limitations of this protocol. The procedure only involves reconnecting the median nerve, while the other 2 nerves are left unrepair. Therefore, the recovery could only yield a portion of the full recovery of the forelimb. The functional recovery of the wrist and individual digits would require modification of the current protocol.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
10-0 Non-Sterile Micro Suture	AROSurgical	TK-107038
4-0 Sharpoint Nylon Black	esutures	A1667N
4-0 Sharpoint Polysyn Taper	esutures	G214N
5 mL syringes	MDC	2045
6-0 silk	Fisher Scientific	NC9742105
ABB-22 V Double Micro Vessel	S&T	00480
B-1 V Single Micro Vessel Clamp	S&T	00462
BD Quincke Spinal needle	esutures	405073
Castroviejo Micro Needle Holders	F.S.T	12061-02
Chatilon DFE II force meter	Ametek	DFE II series
Cotton tiped	JH Supply store	100252
Fine Scissors - Tungsten Carbide	F.S.T	14569-09
Forceps	S&T	FRS-15 RM-8
Gauze Sponges	Fisher Scientific	22-362178
Halsted-Mosquito Hemostats	F.S.T	13009-12
Heparin	JHH Pharmacy	NDC 63323-540-05
I.V. Catheter Radiopaque 24 G	esutures	SMTH5053
Jewelers Bipolar Coagulating Forceps, Stainless Steel	ASSI	103000BPS03
Round Handled Vannas Spring Scissors	F.S.T	15400-12
Special-Forceps	S&T	FRAS-15 RM-8
Vessel Dilators - Balanced Instruments	F.S.T	18602-15