We would like to thank Randall Raish for his excellent videography and editing assistance.

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
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Apply eye lubricant to the mouse&#39;s eyes. Place the mouse on a draped supplemental heating pad to ensure proper body temperature.</li>
	<li>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.</li>
	<li>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).</li>
	<li>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.</li>
	<li>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).</li>
	<li>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).</li>
	<li>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.</li>
	<li>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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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).</li>
</ol>
2. Graft preparation
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Excise the adventitia away from the arterial ends so they have clean edges for anastomosis. 
	&#8203;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.</li>
</ol>
3. Recipient surgery and anastomosis of the vessels
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>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).</li>
	<li>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.</li>
	<li>Dissect the sternomastoid muscle and retract it laterally. Keep the dissected vein protected behind the muscle to avoid direct contact with the retractor.</li>
	<li>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).</li>
	<li>Pass the background material under the external jugular vein and apply the double approximating V3 vessel clamps.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Using a 30 G blunt-tipped needle, flush the inside of the vein with heparinized saline.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Remove the clamps on the venous side. If bleeding is encountered, apply gentle pressure with cotton tips.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Inject 1 mL of warm saline subcutaneously to account for fluid loss during the surgery.</li>
	<li>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.</li>
</ol>

<ol>
	<li>Strome, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laryngeal+transplantation+and+40-month+follow-up.">Laryngeal transplantation and 40-month follow-up.</a> The New England Journal of Medicine. 344 (22), 1676-1679 (2001).</li><li>Hilgers, F. J. M., Ackerstaff, A. H., Aaronson, N. K., Schouwenburg, P. F., Zandwijk, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Physical+and+psychosocial+consequences+of+total+laryngectomy.">Physical and psychosocial consequences of total laryngectomy.</a> Clinical Otolaryngology. 15 (5), 421-425 (1990).</li><li>Heyes, R., Iarocci, A., Tchoukalova, Y., Lott, D. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunomodulatory+role+of+mesenchymal+stem+cell+therapy+in+vascularized+composite+allotransplantation.">Immunomodulatory role of mesenchymal stem cell therapy in vascularized composite allotransplantation.</a> Journal of Transplantation. 2016, (2016).</li><li>Kluyskens, P., Ringoir, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Follow-up+of+a+human+larynx+transplantation.">Follow-up of a human larynx transplantation.</a> Laryngoscope. 80 (8), 1244-1250 (1970).</li><li>Krishnan, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+current+status+of+human+laryngeal+transplantation+in+2017:+A+state+of+the+field+review.">The current status of human laryngeal transplantation in 2017: A state of the field review.</a> Laryngoscope. 127 (8), 1861-1868 (2017).</li><li>Strome, S., Sloman-Moll, E., Wu, J., Samonte, B. R., Strome, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rat+model+for+a+vascularized+laryngeal+allograft.">Rat model for a vascularized laryngeal allograft.</a> Annals of Otology, Rhinology &amp; Laryngology. 101 (11), 950-953 (1992).</li><li>Lorenz, R. R., Dan, O., Nelson, M., Fritz, M. A., Strome, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rat+laryngeal+transplant+model:+technical+advancements+and+a+redefined+rejection+grading+system.">Rat laryngeal transplant model: technical advancements and a redefined rejection grading system.</a> Annals of Otology, Rhinology &amp; Laryngology. 111 (12), 1120-1127 (2002).</li><li>Shipchandler, T. Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+mouse+model+for+studying+laryngeal+transplantation.">New mouse model for studying laryngeal transplantation.</a> Annals of Otology, Rhinology &amp; Laryngology. 118 (6), 465-468 (2009).</li><li>Strome, M., Wu, J., Strome, S., Brodsky, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comparison+of+preservation+techniques+in+a+vascularized+rat+laryngeal+transplant+model.">A comparison of preservation techniques in a vascularized rat laryngeal transplant model.</a> The Laryngoscope. 104 (6), 666-668 (1994).</li><li>Nocini, R., Molteni, G., Mattiuzzi, C., Lippi, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Updates+on+larynx+cancer+epidemiology.">Updates on larynx cancer epidemiology.</a> Chinese Journal of Cancer Research. 32 (1), 18-25 (2020).</li><li>Strome, S., Sloman-Moll, E., Wu, J., Samonte, B. R., Strome, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rat+model+for+a+vascularized+laryngeal+allograft.">Rat model for a vascularized laryngeal allograft.</a> Annals of Otology, Rhinology &amp; Laryngology. 101 (11), (1992).</li><li>Work, W. P., Boles, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Larynx:+Replantation+in+the+dog.">Larynx: Replantation in the dog.</a> Archives of Otolaryngology-Head and Neck Surgery. 82 (4), 401-402 (1965).</li><li>Birchall, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Model+for+experimental+revascularized+laryngeal+allotransplantation.">Model for experimental revascularized laryngeal allotransplantation.</a> British Journal of Surgery. 89 (11), 1470-1475 (2002).</li><li>Nakai, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rat+model+of+laryngeal+transplantation+with+normal+circulation+maintained+by+combination+with+the+tongue.">Rat model of laryngeal transplantation with normal circulation maintained by combination with the tongue.</a> Microsurgery. 23 (2), 135-140 (2003).</li><li>Lott, D. 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The authors declare they have no competing financial interests. Egehan Salepci's travel and living expenses for research were funded by The Scientific and Technological Research Council of Turkey (TUBITAK).

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&#39;s quality of life, and therefore an alternative treatment is desired. Vascularized composite allotransplantation of the larynx can improve a patient&#39;s ability to breathe and speak; however, research is still required before this technique can be utilized c...

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.&#160;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 patients4,5.
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 20026,7. 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.

<table><tbody><tr><td>#1 Paperclips</td><td>Staples</td><td>OP-7404</td><td>Clips are shaped manually to be used as retractors</td></tr><tr><td>1 cc Insulin Syringes&amp;nbsp;</td><td>BD&amp;nbsp;</td><td>329412</td><td>27 G 5/8</td></tr><tr><td>10-0 Ethilon Nylon Suture</td><td>Ethicon</td><td>2870G</td><td></td></tr><tr><td>25 G Precision Glide Needle</td><td>BD&amp;nbsp;</td><td>305125</td><td>1 in</td></tr><tr><td>3 mL Luer-Lok Tip Syringe</td><td>BD&amp;nbsp;</td><td>309657</td><td></td></tr><tr><td>30 G Sterile Standard Blunt Needles</td><td>Cellink</td><td>NZ5300505001</td><td></td></tr><tr><td>5-0 Monocryl Suture</td><td>Ethicon</td><td>Y822G</td><td></td></tr><tr><td>8-0 Ethilon Nylon Suture</td><td>Ethicon</td><td>2815G</td><td></td></tr><tr><td>Adson Forceps</td><td>Fine Science Tools</td><td>11027-12</td><td>Straight, 1 x 2 teeth</td></tr><tr><td>Adventitia scissors</td><td>S&amp;amp;T</td><td>SAS-10</td><td>19 mm, 10 cm, straight</td></tr><tr><td>Angled Forceps</td><td>Fine Science Tools</td><td>00109-11</td><td>45/11 cm</td></tr><tr><td>Artifical Tears Lubricant Opthalmic Ointment</td><td>Akorn Animal Health</td><td>59399-162-35</td><td></td></tr><tr><td>Bandaid Fabric Fingertip</td><td>Cardinal Healthcare</td><td>299399</td><td></td></tr><tr><td>Betadine Solution Swabsticks</td><td>Purdue Products</td><td>67618-153-01</td><td></td></tr><tr><td>Buprenex Injection</td><td>CIII</td><td>12495-0757-1</td><td>0.3 mg/mL</td></tr><tr><td>Clamp applying forceps without lock</td><td>Accurate Surgical &amp;amp; Scientific Instruments</td><td>ASSI.CAF5</td><td>14 cm</td></tr><tr><td>Cotton Swabs</td><td>Puritan</td><td>10806-001-PK</td><td></td></tr><tr><td>DeBakey forceps</td><td></td><td></td><td></td></tr><tr><td>Dermabond Mini</td><td>Cardinal Healthcare</td><td>315999</td><td></td></tr><tr><td>Dissecting Boards</td><td>Mopec</td><td>22-444-314</td><td></td></tr><tr><td>Falcon Sterile Disposable Petri Dish&amp;nbsp;</td><td>Corning</td><td>25373-041</td><td>35 mm</td></tr><tr><td>Fine Scisssors</td><td>Fine Science Tools</td><td>14029-10</td><td>Curved Sharp-Blunt 10 cm</td></tr><tr><td>Golden A5 2-Speed Blade Clipper&amp;nbsp;</td><td>Oster</td><td>008OST-78005-140</td><td>#10</td></tr><tr><td>Hair Remover Sensitive Formula</td><td>Nair</td><td>2260000033</td><td></td></tr><tr><td>Heparin&amp;nbsp;</td><td>Meitheal Pharmaceuticals</td><td>71288-4O2-10</td><td>10,000 USP units per 10 mL</td></tr><tr><td>Isoflurane</td><td>Piramal Healthcare</td><td>66794-013-25</td><td></td></tr><tr><td>Low-Temp Micro Fine Tip Cautery</td><td>Bovie Medical</td><td>AA90</td><td></td></tr><tr><td>Mercian Visibility Background Material</td><td>Synovis Micro Companies</td><td>VB3</td><td>Green</td></tr><tr><td>Microvascular Approximator Clamp without Frame</td><td>Accurate Surgical &amp;amp; Scientific Instruments</td><td>ASSI.ABB11V</td><td>0.4-1 mm Vessel Diameter</td></tr><tr><td>Mouse face mask kit</td><td>Xenotec</td><td>XRK-S</td><td>Small</td></tr><tr><td>Needle holder</td><td>S&amp;amp;T</td><td>C-14 W</td><td>5.5&quot;, 8 mm, 0.4 mm</td></tr><tr><td>Press n&#039; Seal</td><td>Glad</td><td>70441</td><td></td></tr><tr><td>Scalpel</td><td>Braun</td><td>BA210</td><td>10 blade</td></tr><tr><td>Single Mini Vessel Clamp</td><td>Accurate Surgical &amp;amp; Scientific Instruments</td><td>ASSI.ABB11M</td><td>.31 (8 mm), 3 x 1 mm Rnd. Bl., Black Pair</td></tr><tr><td>Stereomicroscope</td><td>Olympus</td><td>SZ61</td><td></td></tr><tr><td>Sterile Alcohol Prep Pads</td><td>Fisherbrand</td><td>06-669-62</td><td></td></tr><tr><td>Sterile Disposable Drape Sheets</td><td>Dynarex</td><td>DYN4410-CASE</td><td></td></tr><tr><td>Sterile Gauze Pads</td><td>Dukal</td><td>1212</td><td></td></tr><tr><td>Sterile Saline&amp;nbsp;</td><td>Hospira</td><td>236173</td><td>NaCl 0.9%</td></tr><tr><td>Sterile Surgical Gloves</td><td>Gammex</td><td>851_A</td><td></td></tr><tr><td>Straight Forceps</td><td>Fine Science Tools</td><td>00108-11</td><td>11 cm</td></tr><tr><td>Tissue forceps</td><td>Accurate Surgical &amp;amp; Scientific Instruments</td><td>ASSI.JFLP3</td><td>13.5 cm, 8 mm, 0.3 mm</td></tr><tr><td>Vannas Pattern Scissors&amp;nbsp;</td><td>Accurate Surgical &amp;amp; Scientific Instruments</td><td>ASSI.SDC15RV</td><td>15 cm, 8 mm, curved 7mm blade</td></tr><tr><td>Vannas Spring Scissors</td><td>Fine Science Tools</td><td>15000-10</td><td>3 mm cutting edge, curved</td></tr><tr><td>Vessel Dilator Tip&amp;nbsp;</td><td>Fine Science Tools</td><td>00126-11</td><td>Diameter 0.1 mm/Angled 10/11 cm</td></tr><tr><td>Vessel Dilator, Classic line</td><td>S&amp;amp;T</td><td>D-5a.3 W</td><td>9 mm, 0.3 mm, angled 10</td></tr></tbody></table>

a heterotopic mouse model for studying laryngeal transplantation

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.

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&#160;4A). If the anastomosis is not effective, the artery will not have pulsation, look partially collapsed, and be pale in color (<st...

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A Heterotopic Mouse Model for Studying Laryngeal Transplantation

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.

Laryngeal heterotopic transplantation, although a technically challenging procedure, offers more scientific analysis and cost benefits compared to other ...

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Research

JoVE Journal

Medicine

2.7K Views.  Mayo Clinic Arizona. Larynxtransplantatiemodellen werden beschreven bij muizen en ratten. Het eerdere werk van Senior alter was gericht op het bedenken van een efficiëntere methode met behulp van de donorcarotisslagaders voor anastomose, waardoor de noodzaak om de superieure schildklierslagaders te ontleden werd vermeden. In deze video beschreven we deze procedure bij muizen.Met dit larynxtransplantatiemodel voor muizen kunnen immunologische studies gemakkelijk en tegen lage kosten worden uitgevoerd. Hoew...

Video: A Heterotopic Mouse Model for Studying Laryngeal Transplantation

Heterotopic Heart Transplantation in Mice

The mouse heterotopic heart transplantation has been used widely since it was introduced by Drs. Corry and Russell in 1973. It is particularly valuable for studying rejection and immune response now that newer transgenic and gene knockout mice are available, and a large number of immunologic reagents have been developed. The heart transplant model is less stringent than the skin transplant models, although technically more challenging. We have developed a modified technique and have completed over 1000 successful cases of heterotopic heart transplantation in mice. When making anastomosis of the ascending aorta and abdominal aorta, two stay sutures are placed at the proximal and distal apexes of recipient abdominal aorta with the donor s ascending aorta, then using 11-0 suture for anastomosis on both side of aorta with continuing sutures. The stay sutures make the anastomosis easier and 11-0 is an ideal suture size to avoid bleeding and thrombosis.
When making anastomosis of pulmonary artery and inferior vena cava, two stay sutures are made at the proximal apex and distal apex of the recipient s inferior vena cava with the donor s pulmonary artery. The left wall of the inferior vena cava and donor s pulmonary artery is closed with continuing sutures in the inside of the inferior vena cava after, one knot with the proximal apex stay suture the right wall of the inferior vena cava and the donor s pulmonary artery are closed with continuing sutures outside the inferior vena cave with 10-0 sutures. This method is easier to perform because anastomosis is made just on the one side of the inferior vena cava and 10-0 sutures is the right size to avoid bleeding and thrombosis. In this article, we provide details of the technique to supplement the video.

The mouse heterotopic heart transplantation model has been proven by many investigators to be an important method for studying mechanisms of rejection and immune response. However, the techniques involved are still challenging. By modifying standard techniques we have had success with more than 1000 transplants.

The mouse heterotopic heart transplantation has been used widely since it was introduced by Drs. Corry and Russell in 1973. It is particularly valuable ...

Biology

Development of Obliterative Bronchiolitis in a Murine Model of Orthotopic Lung Transplantation

Orthotopic lung transplantation in rats was first reported by Asimacopoulos and colleagues in 1971 1. Currently, this method is well accepted and standardized not only for the study of allo-rejection but also between syngeneic strains for examining mechanisms of ischemia-reperfusion injury after lung transplantation. Although the application of the rat and other large animal model 2 contributed significantly to the elucidation of these studies, the scope of those investigations is limited by the scarcity of knockout and transgenic rats. Due to no effective therapies for obliterative bronchiolitis, the leading cause of death in lung transplant patients, there has been an intensive search for pre-clinical models that replicate obliterative bronchiolitis. The tracheal allograft model is the most widely used and may reproduce some of the histopathologic features of obliterative bronchiolitis 3. However, the lack of an intact vasculature with no connection to the recipient's conducting airways, and incomplete pathologic features of obliterative bronchiolitis limit the utility of this model 4. Unlike transplantation of other solid organs, vascularized mouse lung transplants have only recently been reported by Okazaki and colleagues for the first time in 2007 5. Applying the basic principles of the rat lung transplant, our lab initiated the obliterative bronchiolitis model using minor histoincompatible antigen murine orthotopic single-left lung transplants which allows the further study of obliterative bronchiolitis immunopathogenesis6.

Obliterative bronchiolitis is the key impediment to the long-term survival of lung transplant recipients and the lack of a robust preclinical model precludes examining obliterative bronchiolitis immunopathogenesis. Unlike other solid organ transplants, vascularized mouse lung transplantation has only recently been developed. Here we show our independently developed obliterative bronchiolitis model after murine orthotopic single-lung transplantation.

Orthotopic lung transplantation in rats was first reported by Asimacopoulos and colleagues in 1971 1. Currently, this method is well accepted and ...

Heterotopic and Orthotopic Tracheal Transplantation in Mice used as Models to Study the Development of  Obliterative Airway Disease

Obliterative airway disease (OAD) is the major complication after lung transplantations that limits long term survival (1-7).
To study the pathophysiology, treatment and prevention of OAD, different animal models of tracheal transplantation in rodents have been developed (1-7). Here, we use two established models of trachea transplantation, the heterotopic and orthotopic model and demonstrate their advantages and limitations.
For the heterotopic model, the donor trachea is wrapped into the greater omentum of the recipient, whereas the donor trachea is anastomosed by end-to-end anastomosis in the orthotopic model.
In both models, the development of obliterative lesions histological similar to clinical OAD has been demonstrated (1-7).
This video shows how to perform both, the heterotopic as well as the orthotopic tracheal transplantation technique in mice, and compares the time course of OAD development in both models using histology.

This video shows and compares two experimental models to study the development of obliterative airway disease (OAD) in mice, the heterotopic and orthotopic tracheal transplantation model.

Obliterative airway disease (OAD) is the major complication after lung transplantations that limits long term survival (1-7).
To study the ...

Heterotopic Cervical Heart Transplantation in Mice

The heterotopic cervical heart transplantation in mice is a valuable tool in transplant and cardiovascular research. The cuff technique greatly simplifies this model by avoiding challenging suture anastomoses of small vessels thereby reducing warm ischemia time. In comparison to abdominal graft implantation the cervical model is less invasive and the implanted graft is easily accessible for further follow-up examinations. Anastomoses are performed by pulling the ascending aorta of the graft over the cuff with the recipient&#8217;s common carotid artery and by pulling the main pulmonary artery over the cuff with the external jugular vein. Selection of appropriate cuff size and complete mobilization of the vessels are important for successful revascularization. Ischemia-reperfusion (I/R) injury can be minimized by perfusing the graft with a cardioplegic solution and by hypothermia. In this article, we provide technical details for a simplified and improved cuff technique, which should allow surgeons with basic microsurgical skills to perform the procedure with a high success rate.

The cuff technique shortens and facilitates the model of heterotopic cervical heart transplantation in mice by avoiding technically challenging suture anastomoses of small vessels. The technical details shown in this video paper should allow researches to establish this model in their laboratories.

The heterotopic cervical heart transplantation in mice is a valuable tool in transplant and cardiovascular research. The cuff technique greatly simplifies ...

Orthotopic Hind Limb Transplantation in the Mouse

In vivo animal model systems, and in particular mouse models, have evolved into powerful and versatile scientific tools indispensable to basic and translational research in the field of transplantation medicine. A vast array of reagents is available exclusively in this setting, including mono- and polyclonal antibodies for both diagnostic and interventional applications. In addition, a vast number of genotyped, inbred, transgenic, and knock out strains allow detailed investigation of the individual contributions of humoral and cellular components to the complex interplay of an immune response and make the mouse the gold standard for immunological research. 
Vascularized Composite Allotransplantation (VCA) delineates a novel field of transplantation using allografts to replace "like with like" in patients suffering traumatic or congenital tissue loss. This surgical methodological protocol shows the use of a non-suture cuff technique for super-microvascular anastomosis in an orthotopic mouse hind limb transplantation model. The model specifically allows for comparison between established paradigms in solid organ transplantation with a novel form of transplants consisting of various different tissue components. Uniquely, this model allows for the transplantation of a viable vascularized bone marrow compartment and niche that have the potential to exert a beneficial effect on the balance of immune acceptance and rejection. This technique provides a tool to investigate alloantigen recognition and allograft rejection and acceptance, as well as enables the pursuit of functional nerve regeneration studies to further advance this novel field of transplantation.

This novel model for orthotopic hind limb transplantation in the mouse, applying a non-suture cuff technique for super-microvascular anastomosis, provides a powerful tool for in&#160;vivo mechanistic immunological research related to vascularized composite allotransplantation (VCA).

In vivo animal model systems, and in particular mouse models, have evolved into powerful and versatile scientific tools indispensable to basic and ...

Seeding and Implantation of a Biosynthetic Tissue-engineered Tracheal Graft in a Mouse Model

Treatment options for congenital or secondary long segment tracheal defects have historically been limited due to an inability to replace functional tissue. Tissue engineering holds great promise as a potential solution with its ability to integrate cells and signaling molecules into a 3-dimensional scaffold. Recent work with tissue engineered tracheal grafts (TETGs) has seen some success but their translation has been limited by graft stenosis, graft collapse, and delayed epithelialization. In order to investigate the mechanisms driving these issues, we have developed a mouse model for tissue engineered tracheal graft implantation. TETGs were constructed using electrospun polymers polyethylene terephthalate (PET) and polyurethane (PU) in a mixture of PET and PU (20:80 percent weight). Scaffolds were then seeded using bone marrow mononuclear cells isolated from 6-8 week-old C57BL/6 mice by gradient centrifugation. Ten million cells per graft were seeded onto the lumen of the scaffold and allowed to incubate overnight before implantation between the third and seventh tracheal rings. These grafts were able to recapitulate the findings of stenosis and delayed epithelialization as demonstrated by histological analysis and lack of Keratin 5 and Keratin 14 basal epithelial cells on immunofluorescence. This model will serve as a tool for investigating cellular and molecular mechanisms involved in host remodeling.

Graft stenosis poses a critical obstacle in tissue engineered airway replacement. To investigate cellular mechanisms underlying stenosis, we utilize a murine model of tissue engineered tracheal replacement with seeded bone marrow mononuclear cells (BM-MNC). Here, we detail our protocol, including scaffold manufacturing, BM-MNC isolation, graft seeding, and implantation.

Treatment options for congenital or secondary long segment tracheal defects have historically been limited due to an inability to replace functional ...

Bioengineering

Mouse Heterotopic Cervical Cardiac Transplantation Utilizing Vascular Cuffs

Murine models of cardiac transplantation are frequently utilized to study ischemia-reperfusion injury, innate and adaptive immune responses after transplantation, and the impact of immunomodulatory therapies on graft rejection. Heterotopic cervical heart transplantation in mice was first described in 1991 using sutured anastomoses and subsequently modified to include cuff techniques. This modification allowed for improved success rates, and since then, there have been multiple reports that have proposed further technical improvements. However, translation into more widespread utilization remains limited due to the technical difficulty associated with graft anastomoses, which requires precision to achieve adequate length and caliber of the cuffs to avoid vascular anastomotic twisting or excessive tension, which can result in damage to the graft. The present protocol describes a modified technique for performing heterotopic cervical cardiac transplantation in mice which involves cuff placement on the recipient's common carotid artery and the donor's pulmonary artery in alignment with the direction of the blood flow.

Mouse cardiac transplantation models represent valuable research tools for studying transplantation immunology. The present protocol details mouse heterotopic cervical cardiac transplantation that involves the placement of cuffs on the recipient's common carotid artery and the donor's pulmonary artery trunk to allow for laminar blood flow.

Murine models of cardiac transplantation are frequently utilized to study ischemia-reperfusion injury, innate and adaptive immune responses after ...

Immunology and Infection

A Modified Cuff Technique for Mouse Cervical Heterotopic Heart Transplantation Model

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&#39; long-term survival treated with cardiac allografts. In a mouse model, abdominal donor heart implantation is commonly performed with an end-to-end&#160;anastomosis to the recipient&#39;s aorta and inferior vena cava using stitches. In this model, the donor&#39;s heart is implanted by end-to-end anastomosis to the recipient&#39;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.

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.

Cardiac allograft rejection limits the long-term survival of patients after heart transplantation. A mouse heart transplantation model is ideal for ...

Exploring Obliterative Airway Disease in Lung Transplants

Murine Intrapulmonary Tracheal Transplantation: A Model for Investigating Obliterative Airway Disease After Lung Transplantation

Murine intrapulmonary tracheal transplantation (IPTT) is used as a model of obliterative airway disease (OAD) following lung transplantation. Initially reported by our team, this model has gained use in the study of OAD due to its high technical reproducibility and suitability for investigating immunological behaviors and therapeutic interventions.
In the IPTT model, a rodent tracheal graft is directly inserted into the recipient's lung through the pleura. This model is distinct from the heterotopic tracheal transplantation (HTT) model, wherein grafts are transplanted into subcutaneous or omental sites, and from the orthotopic tracheal transplantation (OTT) model in which the donor trachea replaces the recipient's trachea. 
Successful implementation of the IPTT model requires advanced anesthetic and surgical skills. Anesthetic skills include endotracheal intubation of the recipient, setting appropriate ventilatory parameters, and appropriately timed extubation after recovery from anesthesia. Surgical skills are essential for precise graft placement within the lung and for ensuring effective sealing of the visceral pleura to prevent air leakage and bleeding. In general, the learning process takes approximately 2 months.
In contrast to the HTT and OTT models, in the IPTT model, the allograft airway develops airway obliteration in the relevant lung microenvironment. This allows investigators to study lung-specific immunological and angiogenic processes involved in airway obliteration after lung transplantation. Furthermore, this model is also unique in that it exhibits tertiary lymphoid organs (TLOs), which are also seen in human lung allografts. TLOs are comprised of T and B cell populations and characterized by the presence of high endothelial venules that direct immune cell recruitment; therefore, they are likely to play a crucial role in graft acceptance and rejection. We conclude that the IPTT model is a useful tool for studying intrapulmonary immune and profibrotic pathways involved in the development of airway obliteration in the lung transplant allograft.

Author Spotlight: Investigating the Key Factors of Obliterative Bronchiolitis After Lung Transplantation

The murine intrapulmonary tracheal transplantation (IPTT) model is valuable for studying obliterative airway disease (OAD) after lung transplantation. It offers insights into lung-specific immunological and angiogenic behavior in airway obliteration after allotransplantation with high reproducibility. Here, we describe the IPTT procedure and its expected results.

Murine intrapulmonary tracheal transplantation (IPTT) is used as a model of obliterative airway disease (OAD) following lung transplantation. Initially ...

A Simple Cuff Technique for Murine Left Lung Transplantation

Over the past decade, our laboratory has made significant progress in developing and refining vascularized mouse lung transplantation models using an efficient and highly reliable "cuff technique" of transplantation. This article describes a sophisticated and comprehensive method for orthotopic lung transplantation in a vascularized orthotopic lung model, representing the most physiologic and clinically relevant model of mouse lung transplantation to date. The transplantation process consists of two distinct stages: donor harvest and subsequent implantation into the recipient. The method has been successfully mastered, and with several months of sufficient training, a skilled practitioner can perform the procedure in approximately 90 min from skin-to-skin. Surprisingly, once individuals overcome the initial learning curve, the survival rate during the perioperative period approaches nearly 100%. The mouse model allows for the use of multiple commercially available transgenic and mutant strains of mice, enabling the study of tolerance and rejection. Additionally, the unique features of this model make it a valuable tool for investigating tumor biology and immunology.

The present protocol describes a cuff technique for a mouse left lung transplantation model. This technique has been developed over several years and has performed well, serving effectively in immunological research.

Over the past decade, our laboratory has made significant progress in developing and refining vascularized mouse lung transplantation models using an ...

A Heterotopic Mouse Model for Studying Laryngeal Transplantation

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