A Heterotopic Mouse Model for Studying Laryngeal Transplantation

Summary

The aim of this manuscript is to describe the microsurgical steps required to perform a heterotopic laryngeal transplant in mice. The advantages of this mouse model compared to other animal models of laryngeal transplantation are its cost-effectiveness and the availability of immunologic assays and data.

Abstract

Laryngeal heterotopic transplantation, although a technically challenging procedure, offers more scientific analysis and cost benefits compared to other animal models. Although first described by Shipchandler et al. in 2009, this technique is not widely used, possibly due to the difficulties in learning the microsurgical technique and time required to master it. This paper describes the surgical steps in detail, as well as potential pitfalls to avoid, in order to encourage effective use of this technique.

In this model, the bilateral carotid arteries of the donor larynx are anastomosed to the recipient carotid artery and external jugular vein, allowing for blood flow through the graft. Blood flow can be confirmed intraoperatively by the visualization of blood filling in the graft bilateral carotid arteries, reddening of the thyroid glands of the graft, and bleeding from micro vessels in the graft. The crucial elements for success include delicate preservation of the graft vessels, making the correct size arteriotomy and venotomy, and using the appropriate number of sutures on the arterial-arterial and arterial-venous anastomoses to secure vessels without leakage and prevent occlusion.

Anyone can become proficient in this model with sufficient training and perform the procedure in approximately 3 h. If performed successfully, this model allows for immunologic studies to be performed with ease and at low cost.

Introduction

For patients suffering from irreparable laryngeal damage or laryngeal cancer, a total laryngectomy is often the only option¹. Total laryngectomy leaves patients without the ability to breathe and speak on their own, in addition to experiencing social and psychological distress². Patients with laryngeal cancer who need a total laryngectomy are excellent potential candidates for laryngeal transplantation. While human laryngeal transplantation in the setting of irreparable laryngeal damage has been performed, allotransplantation of the larynx is currently avoided in these patients due to the fear of tumor recurrence, the possibility of chronic rejection, and donor-derived infections³. Immunosuppression is the primary cause of these concerns. The dramatic loss of the first partial laryngeal transplant patient due to tumor recurrence after conventional immunosuppressive treatment is evidence that an appropriate immunosuppressive regimen should be devised before further attempts are made for transplantation in laryngeal cancer patients^4,5.

To better understand the host immune response to a transplanted larynx, the first laryngeal transplant model in rats was developed in 1992 by Strome, and improvements to the surgical technique were made in 2002^6,7. Although this model is effective for studying laryngeal transplantation, the lack of rat-specific immunological agents and the higher cost associated with rat models led to the development of a new mouse model for studying laryngeal transplantation in 2009⁸.

The main application of the described technique is to study different immunosuppressive drug regimens in laryngeal transplantation. Improving current immunosuppressive therapies may broaden the candidate pool and lead to safe transplantation in cancer patients. The benefits of this mouse model are its cost-effectiveness and the wide availability of immunologic data and reagents.

Teams working on immunosuppressive treatment regimens for laryngeal transplantation can use this method to collect a large volume of immunologic data, and different drug regimens can be rapidly tested and compared. Other potential treatment modalities that can modulate the immune response to the transplant, such as stem cell injections, can also be tested using this model. Finally, experiments can be devised to observe long-term systemic effects of laryngeal transplantation by extending the follow-up period.

The technique described here uses end-to-side anastomoses to provide arterial and venous flow to a heterotopic larynx graft. The graft is a laryngotracheoesophageal (LTE) complex comprising the larynx, thyroid glands, parathyroid glands, trachea, and esophagus of the donor, with bilateral carotid arteries and pedicles intact. One donor carotid artery is anastomosed to the recipient carotid artery and provides arterial blood flow, while the other donor carotid artery is anastomosed to the recipient external jugular vein and provides venous blood flow (Figure 1).

Several modifications were made to the surgical technique of the rat model to ensure success in the mouse model. For instance, an inhaled anesthetic agent was used instead of an injectable agent to increase control over the depth of anesthesia and reduce complications. Continuous suturing is used in the arterial-arterial anastomosis in rats; however, due to the smaller size of mouse vessels, this is technically difficult and can cause narrowing of the vessel lumen⁷. As a result, interrupted sutures are used in the mouse model and result in improved vessel patency. Additionally, in the rat model, the superior thyroid artery (STA) pedicle is dissected out and visualized. Given the smaller size of the STA in mice, this dissection could result in damage to and even transection of the STA. As a result, it is not dissected in the mouse model. Instead, nearby fascia is preserved to ensure that the STA is kept intact.

The major potential pitfalls of this technique include damaging the donor LTE complex pedicles, making an incorrectly sized arteriotomy or venotomy, vessel occlusion at the anastomosis sites, or leaving gaps at the anastomosis sites which may cause bleeding. To avoid these missteps, care must be taken when procuring the donor graft by leaving a cuff of tissue around the STA pedicle. The arteriotomy and venotomy should be large enough to allow blood flow but small enough to prevent leakage. An appropriate number of sutures should be used for the anastomoses to close any gaps, but not too many to occlude the vessels.

If familiarity with the microsurgical techniques is obtained, this procedure can be performed in approximately 3 h. This laryngeal transplantation model can be performed reliably in mice and used to study the host immune response after vascularized composite allotransplantation.

Protocol

This research was performed in compliance with the Mayo Clinic Institutional Animal Care and Use Committee (IACUC). BalbC mice (10-12 weeks old) were used as donors and C57/BL6 mice (10-12 weeks old) were used as recipients because their major histocompatibility complexes, H-2Db and H-2Kb, respectively, are immunologically incompatible, and therefore the immune response to the graft can be further studied. All instruments used during surgery were sterilized (see Supplemental Figure S1 and Supplemental Figure S2), and the surgical field was kept sterile throughout the protocol per IACUC instructions.

1. Donor surgery and graft procurement

1. Inject the donor with extended-release buprenorphine (3.25 mg/kg body weight subcutaneously) 30 min before starting the procedure. Place the mouse in the rodent anesthesia box for anesthesia induction at 3% isoflurane delivered with 1 L/min O₂ flow. After the animal is fully anesthetized, transfer the mouse to the shaving area and administer 1.5% isoflurane with 1 L/min O₂ flow for maintenance of anesthesia. Confirm the depth of anesthesia with a toe pinch.
2. Shave the chest and neck of the mouse up to the jawline and apply depilatory cream. After 30 s, wipe the cream off with a sterile gauze pad wetted with water and transfer the mouse to the surgical area.
3. Apply eye lubricant to the mouse's eyes. Place the mouse on a draped supplemental heating pad to ensure proper body temperature.
4. Immobilize the mouse, prep the surgical area thrice with povidone iodine and alcohol, and then drape the mouse.
   NOTE: Check the depth of anesthesia by a toe pinch and maintain anesthesia at 1.5% isoflurane and 1 L/min O₂ flow via a face mask throughout the procedure.
5. Make a small horizontal incision just superior to the suprasternal notch. Using fine scissors, elevate the skin bilaterally through that incision up to the mandible. Excise a trapezoid-shaped skin segment resulting in exposure of the bilateral salivary glands, sternomastoid muscles, digastric muscles, and superior aspect of the sternum (Figure 2A).
6. Excise the bilateral salivary glands using cautery at the superior part where a small vein traveling through the gland is visualized (Figure 2A).
   NOTE: If care is not taken to cauterize the vessel before removing the gland, significant bleeding can be encountered at this step.
7. Gently retract the lymphoid and adipose tissues laterally to expose the sternomastoid and strap muscles. Dissect the bilateral sternomastoid muscles from the surrounding tissues and retract them laterally using retractors.
   NOTE: This maneuver will fully expose the LTE complex with strap muscles and provide a comfortable working space for dissection of the carotid arteries (Figure 2B).
8. Make a midline incision between the strap muscles and bilaterally excise them, taking care not to damage the underlying thyroid gland and leaving it on the LTE complex.
   NOTE: After this step, the bilateral carotid arteries should be visible (Figure 2C).
9. Circumferentially dissect the common carotid arteries to the level of the clavicle inferiorly and to the level of the carotid bifurcation superiorly. Dissect the vagus nerve and the internal jugular vein from the carotid arteries. Do not include them in the procured graft.
   NOTE: The superior thyroid artery can be seen just superior to the bifurcation traveling medially. Leave the thin fascia surrounding this vessel intact and do not attempt to dissect it circumferentially. This vessel supplies the blood flow to the LTE complex and will serve as the pedicle after transplantation. Preservation of this vessel is of utmost importance.
10. Dissect the internal and external carotid arteries far enough superiorly to be able to ligate and divide. If there is difficulty with visualization of the vessels, use a separate retractor to retract the digastric muscles laterally.
    NOTE: Avoid dissecting the external carotid artery closer to the bifurcation to prevent any damage to the superior thyroid artery. At this step, the occipital artery, which branches from the external carotid and follows parallel to the internal carotid artery, can be encountered and should not be confused with the internal carotid artery, which is larger and found deeper (Figure 2D).
11. Using 8-0 nylon sutures, ligate the internal carotid arteries 2 to 3 mm superior to the carotid bifurcation. Ligate the external carotid arteries at least 3 mm superior to the branching point of the superior thyroid artery.
    NOTE: After these ligations, the LTE complex will be pedicled via the superior thyroid arteries to the bilateral carotid arteries.
12. Ligate the common carotid arteries at the level of the sternum and cut all the ligated vessels bilaterally. Keep the vascular pedicles on top of the LTE complex to avoid accidental damage during further dissection. To prevent any gas leakage or inadvertent loss of anesthesia, make sure the animal has expired before making any airway cuts.
13. Divide the infrahyoid muscles at the level of the hyoid. Create an anterior pharyngotomy just inferior to the hyoid. Carry the incision down to the prevertebral fascia to free the LTE complex superiorly.
14. Transect the trachea below the fifth tracheal ring and carry the incision through the esophagus down to the prevertebral fascia to free the LTE complex inferiorly. Free the trachea and esophagus from the underlying prevertebral fascia from an inferior to superior direction.
    NOTE: Keep the vascular pedicles on top of the LTE complex to avoid accidental damage during further dissection.
15. Create an anterior pharyngotomy just inferior to the hyoid. Carry the incision down to the prevertebral fascia to free the LTE complex superiorly. Divide any remaining lymphoid or connective tissue attachments between the LTE complex and the surrounding tissue. Remove the graft.
    NOTE: The graft contains the donor larynx, trachea, thyroid glands, parathyroid glands, esophagus, and laryngeal muscles as a composite unit (Figure 2E).

2. Graft preparation

1. Place the procured graft in a sterile Petri dish and wash it with normal saline to get rid of any blood clots. Using micro forceps, gently milk the blood and clots out of the bilateral carotid arteries. Dilate the bilateral carotid arteries using 1 mm microdilators.
2. Using a 30 G blunt-tipped needle, inject approximately 2 mL of heparinized saline in each carotid artery to flush the graft.
   NOTE: Blood and saline can be seen flushing out of the contralateral carotid artery and small free vessel ends, which confirms intact superior thyroid arteries.
3. Excise the adventitia away from the arterial ends so they have clean edges for anastomosis.
   ​NOTE: The graft can be left in heparinized saline and a short break can be taken for up to 3 h before transplanting it into the recipient⁹.

3. Recipient surgery and anastomosis of the vessels

1. Prepare the recipient mouse in the same manner as described for the donor following the anesthesia induction, shaving, and surgical preparation steps. Inject the recipient mouse with extended-release analgesic subcutaneously 30 min prior to the start of surgery.
2. Using a scalpel, make a midline neck incision extending from the jawline superiorly to the sternum inferiorly. Elevate the skin on the left side and retract it laterally.
3. Excise the left salivary gland, cauterizing the superior vessels as previously described. Excise the adipose and lymphoid tissue by dividing any visible vessels with low-temperature cautery, taking care not to damage the underlying external jugular vein (Figure 3A).
4. Dissect the external jugular vein circumferentially. Use at least 5 mm of clear length of the vessel for anastomosis. Use low-temperature cautery or ligate and divide any relatively large veins branching from the jugular vein.
5. Dissect the sternomastoid muscle and retract it laterally. Keep the dissected vein protected behind the muscle to avoid direct contact with the retractor.
6. Excise the left strap muscles to gain access to the recipient carotid artery. Circumferentially dissect the common carotid artery from the clavicle inferiorly up to the carotid bifurcation superiorly (Figure 3B).
7. Pass the background material under the external jugular vein and apply the double approximating V3 vessel clamps.
8. Place a 10-0 nylon suture through the anterior wall of the external jugular vein at the location of the desired venotomy and use this suture to pull anteriorly and tent the vessel.
9. Cut to the suture with microscissors just deep enough to create the proper size single-slit venotomy and ensure the cut is completely through the venous wall.
10. Using a 30 G blunt-tipped needle, flush the inside of the vein with heparinized saline.
11. Place the donor LTE complex between the recipient left carotid artery and the left external jugular vein. Align the free end of the donor left carotid artery toward the recipient left external jugular vein and bevel the ends of the vessels with sharp scissors.
12. Using four 10-0 nylon interrupted sutures, anastomose the donor left carotid artery and recipient left external jugular vein in an end-to-side fashion.
13. Slide the background material under the recipient common carotid artery and place the double approximating A3 vessel clamps on the recipient common carotid artery. Create an arteriotomy in the same fashion as the venotomy.
    NOTE: Make sure the arteriotomy is the same size as the lumen of the donor carotid artery. If it is too large, profuse bleeding will occur after the clamps are removed. If it is too small, blood flow to the graft will be obstructed.
14. Anastomose the donor right carotid artery to the recipient left carotid artery in an end-to-side fashion using six 10-0 nylon interrupted sutures.
    NOTE: Correct microvascular technique should be respected throughout the vessel anastomosis. Passing through the backwall results in considerably constricted blood flow, endangering survival of the graft. Due to the small size of the vessels, trying to redo the sutures is very difficult.
15. Remove the clamps on the venous side. If bleeding is encountered, apply gentle pressure with cotton tips.
16. Remove the clamps on the artery and immediately apply gentle pressure with cotton tips.
    NOTE: Some bleeding is expected at this step, which usually stops after 1 min with gentle pressure.
17. Check the integrity of blood flow in the artery and the vein.
    NOTE: With intact arterial flow, pulsation of the donor carotid artery is usually seen, and the donor thyroid gland changes from its flushed transparent color back to its original reddish color. Red coloration of the small vessels on the LTE complex can also be observed.
18. Irrigate the surgical field with heparinized saline and close the skin incision with a 5-0 monofilament suture in a running fashion. Apply antibiotic ointment or a skin adhesive on the incision.
19. Inject 1 mL of warm saline subcutaneously to account for fluid loss during the surgery.
20. Stop the anesthesia and transfer the mouse to a recovery cage. Observe the mouse on a heating pad until it is fully awake to avoid hypothermia.

Results

Confirmation of successful transplantation
Using the protocol described above, it is possible to assess blood flow to the LTE complex by observing the pulsation of the donor carotid artery after removing the vessel clamps. Pulsation is typically visible, and immediate red coloration of the donor artery confirms active blood flow (Figure 4A). If the anastomosis is not effective, the artery will not have pulsation, look partially collapsed, and be pale in color (

Discussion

The incidence and prevalence of laryngeal cancer have increased by 12% and 24%, respectively, during the last three decades, and many of these patients undergo a laryngectomy for treatment¹⁰. This procedure significantly worsens a person's quality of life, and therefore an alternative treatment is desired. Vascularized composite allotransplantation of the larynx can improve a patient's ability to breathe and speak; however, research is still required before this technique can be utilized c...

Materials

Name	Company	Catalog Number	Comments
#1 Paperclips	Staples	OP-7404	Clips are shaped manually to be used as retractors
1 cc Insulin Syringes	BD	329412	27 G 5/8
10-0 Ethilon Nylon Suture	Ethicon	2870G
25 G Precision Glide Needle	BD	305125	1 in
3 mL Luer-Lok Tip Syringe	BD	309657
30 G Sterile Standard Blunt Needles	Cellink	NZ5300505001
5-0 Monocryl Suture	Ethicon	Y822G
8-0 Ethilon Nylon Suture	Ethicon	2815G
Adson Forceps	Fine Science Tools	11027-12	Straight, 1 x 2 teeth
Adventitia scissors	S&T	SAS-10	19 mm, 10 cm, straight
Angled Forceps	Fine Science Tools	00109-11	45/11 cm
Artifical Tears Lubricant Opthalmic Ointment	Akorn Animal Health	59399-162-35
Bandaid Fabric Fingertip	Cardinal Healthcare	299399
Betadine Solution Swabsticks	Purdue Products	67618-153-01
Buprenex Injection	CIII	12495-0757-1	0.3 mg/mL
Clamp applying forceps without lock	Accurate Surgical & Scientific Instruments	ASSI.CAF5	14 cm
Cotton Swabs	Puritan	10806-001-PK
DeBakey forceps
Dermabond Mini	Cardinal Healthcare	315999
Dissecting Boards	Mopec	22-444-314
Falcon Sterile Disposable Petri Dish	Corning	25373-041	35 mm
Fine Scisssors	Fine Science Tools	14029-10	Curved Sharp-Blunt 10 cm
Golden A5 2-Speed Blade Clipper	Oster	008OST-78005-140	#10
Hair Remover Sensitive Formula	Nair	2260000033
Heparin	Meitheal Pharmaceuticals	71288-4O2-10	10,000 USP units per 10 mL
Isoflurane	Piramal Healthcare	66794-013-25
Low-Temp Micro Fine Tip Cautery	Bovie Medical	AA90
Mercian Visibility Background Material	Synovis Micro Companies	VB3	Green
Microvascular Approximator Clamp without Frame	Accurate Surgical & Scientific Instruments	ASSI.ABB11V	0.4-1 mm Vessel Diameter
Mouse face mask kit	Xenotec	XRK-S	Small
Needle holder	S&T	C-14 W	5.5", 8 mm, 0.4 mm
Press n' Seal	Glad	70441
Scalpel	Braun	BA210	10 blade
Single Mini Vessel Clamp	Accurate Surgical & Scientific Instruments	ASSI.ABB11M	.31 (8 mm), 3 x 1 mm Rnd. Bl., Black Pair
Stereomicroscope	Olympus	SZ61
Sterile Alcohol Prep Pads	Fisherbrand	06-669-62
Sterile Disposable Drape Sheets	Dynarex	DYN4410-CASE
Sterile Gauze Pads	Dukal	1212
Sterile Saline	Hospira	236173	NaCl 0.9%
Sterile Surgical Gloves	Gammex	851_A
Straight Forceps	Fine Science Tools	00108-11	11 cm
Tissue forceps	Accurate Surgical & Scientific Instruments	ASSI.JFLP3	13.5 cm, 8 mm, 0.3 mm
Vannas Pattern Scissors	Accurate Surgical & Scientific Instruments	ASSI.SDC15RV	15 cm, 8 mm, curved 7mm blade
Vannas Spring Scissors	Fine Science Tools	15000-10	3 mm cutting edge, curved
Vessel Dilator Tip	Fine Science Tools	00126-11	Diameter 0.1 mm/Angled 10/11 cm
Vessel Dilator, Classic line	S&T	D-5a.3 W	9 mm, 0.3 mm, angled 10