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. G., Dan, O., Lu, L., 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=Long-term+laryngeal+allograft+survival+using+low-dose+everolimus.">Long-term laryngeal allograft survival using low-dose everolimus.</a> Otolaryngology-Head and Neck Surgery. 142 (1), 72-78 (2010).</li><li>Lott, D. G., Russell, J. O., Khariwala, S. S., Dan, O., 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=Ten-month+laryngeal+allograft+survival+with+use+of+pulsed+everolimus+and+anti-&#945;&#946;+T-cell+receptor+antibody+immunosuppression.">Ten-month laryngeal allograft survival with use of pulsed everolimus and anti-&#945;&#946; T-cell receptor antibody immunosuppression.</a> Annals of Otology, Rhinology &amp; Laryngology. 120 (2), 131-136 (2011).</li><li>Lott, D. G., Dan, O., Lu, L., 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=Decoy+NF-&#954;B+fortified+immature+dendritic+cells+maintain+laryngeal+allograft+integrity+and+provide+enhancement+of+regulatory+T+cells.">Decoy NF-&#954;B fortified immature dendritic cells maintain laryngeal allograft integrity and provide enhancement of regulatory T cells.</a> The Laryngoscope. 120 (1), 44-52 (2010).</li></ol>

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.

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	<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&#160;</td>
			<td>BD&#160;</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&#160;</td>
			<td>305125</td>
			<td>1 in</td>
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		<tr>
			<td>3 mL Luer-Lok Tip Syringe</td>
			<td>BD&#160;</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&#38;T</td>
			<td>SAS-10</td>
			<td>19 mm, 10 cm, straight</td>
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		<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>
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		<tr>
			<td>Clamp applying forceps without lock</td>
			<td>Accurate Surgical &#38; 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&#160;</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&#160;</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&#160;</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 &#38; 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&#38;T</td>
			<td>C-14 W</td>
			<td>5.5&#34;, 8 mm, 0.4 mm</td>
		</tr>
		<tr>
			<td>Press n&#39; 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 &#38; 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&#160;</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 &#38; Scientific Instruments</td>
			<td>ASSI.JFLP3</td>
			<td>13.5 cm, 8 mm, 0.3 mm</td>
		</tr>
		<tr>
			<td>Vannas Pattern Scissors&#160;</td>
			<td>Accurate Surgical &#38; 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&#160;</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&#38;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...

Watch this Scientific Journal Video about A Heterotopic Mouse Model for Studying Laryngeal Transplantation at JoVE.com

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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2.7K Views.  Mayo Clinic Arizona. 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.

Video: A Heterotopic Mouse Model for Studying Laryngeal Transplantation

A Mouse Model of in Utero Transplantation

The transplantation of stem cells and viruses in utero has tremendous potential for treating congenital disorders in the human fetus. For example, in utero transplantation (IUT) of hematopoietic stem cells has been used to successfully treat patients with severe combined immunodeficiency.1,2 In several other conditions, however, IUT has been attempted without success.3 Given these mixed results, the availability of an efficient non-human model to study the biological sequelae of stem cell transplantation and gene therapy is critical to advance this field. We and others have used the mouse model of IUT to study factors affecting successful engraftment of in utero transplanted hematopoietic stem cells in both wild-type mice4-7 and those with genetic diseases.8,9 The fetal environment also offers considerable advantages for the success of in utero gene therapy. For example, the delivery of adenoviral10, adeno-associated viral10, retroviral11, and lentiviral vectors12,13 into the fetus has resulted in the transduction of multiple organs distant from the site of injection with long-term gene expression. in utero gene therapy may therefore be considered as a possible treatment strategy for single gene disorders such as muscular dystrophy or cystic fibrosis. Another potential advantage of IUT is the ability to induce immune tolerance to a specific antigen. As seen in mice with hemophilia, the introduction of Factor IX early in development results in tolerance to this protein.14 
In addition to its use in investigating potential human therapies, the mouse model of IUT can be a powerful tool to study basic questions in developmental and stem cell biology. For example, one can deliver various small molecules to induce or inhibit specific gene expression at defined gestational stages and manipulate developmental pathways. The impact of these alterations can be assessed at various timepoints after the initial transplantation. Furthermore, one can transplant pluripotent or lineage specific progenitor cells into the fetal environment to study stem cell differentiation in a non-irradiated and unperturbed host environment.
The mouse model of IUT has already provided numerous insights within the fields of immunology, and developmental and stem cell biology. In this video-based protocol, we describe a step-by-step approach to performing IUT in mouse fetuses and outline the critical steps and potential pitfalls of this technique.

A Mouse Model of in Utero Transplantation

The mouse model of in utero transplantation is a versatile tool that can be used to study the potential clinical applications of stem cell transplantation and gene therapy in the fetus. In this protocol, we present a general approach to performing this technique

The transplantation of stem cells and viruses in utero has tremendous potential for treating congenital disorders in the human fetus.  For example, in ...

A Modified Method for Heterotopic Mouse Heart Transplantion

Mice are often used as heart transplant donors and recipients in studies of transplant immunology due to the wide range of transgenic mice and reagents available. A difficulty is presented due to the small size of the animal and the considerable technical challenges of the microsurgery involved in heart transplantation. In particular, a high rate of technical failure early after transplantation may result from recipient death and post-operative complications such as hind limb paralysis or a non-beating heart. Here, the complete technique for heterotopic mouse heart transplantation is demonstrated, involving harvesting the donor heart and its subsequent implantation into a recipient mouse. The donor heart is harvested immediately following in situ perfusion with cold heparinized saline and transection of the ascending aorta and pulmonary artery. The recipient operation involves preparation of the abdominal aorta and inferior vena cava (IVC), followed by end-to-side anastomosis of the donor aorta with the recipient aorta using a single running 10-0 microsuture and a similar anastomosis of the donor pulmonary artery with the recipient IVC. Following the operation the animal is injected with 0.6 ml normal saline subcutaneously and allowed to recover on a 37&#160;&#176;C heating pad. The results from 227 mouse heart transplants are summarized with a success rate at 48 hr of 86.8%. Of the 13.2% failures within 48 hr, 5 (2.2%) experienced hind limb paralysis, 10 (4.4%) had a non-beating heart due to graft ischemic injury and/or thrombosis, while 15 (6.6%) died within 48 hr.

A technique is demonstrated for the microsurgical procedure for heterotopic transplantation of hearts in mice, including simplified methods for donor harvesting and recipient vessel anastomosis.

Mice are often used as heart transplant donors and recipients in studies of transplant immunology due to the wide range of transgenic mice and reagents ...

Murine Heterotopic Heart Transplant Technique

It is now over forty years since this technique was first reported by Corry, Wynn and Russell. Although it took some years for other labs to become proficient in and utilize this technique, it is now widely used by many laboratories around the world. A significant refinement to the original technique was developed and reported in 2001 by Niimi. Described here are the techniques that have evolved over more than a decade in the hands of three surgeons (Plenter, Grazia, Pietra) in our center. These techniques are now being passed on to a younger generation of surgeons and researchers.
Based largely on the Niimi experience, the procedures used have evolved in the fine details - details which we will endeavor to relate here in such a way that others may be able to use this very useful model. Like Niimi, we have found that a video aid to learning is a priceless resource for the beginner.

The manuscript describes the steps required to perform the heterotopic heart transplant in the mouse.

It is now over forty years since this technique was first reported by Corry, Wynn and Russell. Although it took some years for other labs to become ...

Transplantation of Pulmonary Valve Using a Mouse Model of Heterotopic Heart Transplantation

Tissue engineered heart valves, especially decellularized valves, are starting to gain momentum in clinical use of reconstructive surgery with mixed results. However, the cellular and molecular mechanisms of the neotissue development, valve thickening, and stenosis development are not researched extensively. To answer the above questions, we developed a murine heterotopic heart valve transplantation model. A heart valve was harvested from a valve donor mouse and transplanted to a heart donor mouse. The heart with a new valve was transplanted heterotopically to a recipient mouse. The transplanted heart showed its own heartbeat, independent of the recipient&#8217;s heartbeat. The blood flow was quantified using a high frequency ultrasound system with a pulsed wave Doppler. The flow through the implanted pulmonary valve showed forward flow with minimal regurgitation and the peak flow was close to 100 mm/sec. This murine model of heart valve transplantation is highly versatile, so it can be modified and adapted to provide different hemodynamic environments and/or can be used with various transgenic mice to study neotissue development in a tissue engineered heart valve.

In order to understand the cellular and molecular mechanisms underlying neotissue formation and stenosis development in tissue engineered heart valves, a murine model of heterotopic heart valve transplantation was developed. A pulmonary heart valve was transplanted to recipient using the heterotopic heart transplantation technique.

Tissue engineered heart valves, especially decellularized valves, are starting to gain momentum in clinical use of reconstructive surgery with mixed ...

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 ...

A Novel Microsurgical Model for Heterotopic, En Bloc Chest Wall, Thymus, and Heart Transplantation in Mice

Exploration of novel strategies in organ transplantation to prolong allograft survival and minimizing the need for long-term maintenance immunosuppression must be pursued. Employing vascularized bone marrow transplantation and co-transplantation of the thymus have shown promise in this regard in various animal models.1-11 Vascularized bone marrow transplantation allows for the uninterrupted transfer of donor bone marrow cells within the preserved donor microenvironment, and the incorporation of thymus tissue with vascularized bone marrow transplantation has shown to increase T-cell chimerism ultimately playing a supportive role in the induction of immune regulation. The combination of solid organ and vascularized composite allotransplantation can uniquely combine these strategies in the form of a novel transplant model. Murine models serve as an excellent paradigm to explore the mechanisms of acute and chronic rejection, chimerism, and tolerance induction, thus providing the foundation to propagate superior allograft survival strategies for larger animal models and future clinical application. Herein, we developed a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique. The experience in syngeneic and allogeneic transplant settings is described for future broader immunological investigations via an instructional manuscript and video supplement.

To study combined solid organ and vascularized composite allotransplantation, we describe a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique.

Exploration of novel strategies in organ transplantation to prolong allograft survival and minimizing the need for long-term maintenance immunosuppression ...

Mouse Model for Pancreas Transplantation Using a Modified Cuff Technique

Mouse models have several advantages in transplantation research, including easy handling, a variety of genetically well-defined strains, and the availability of the widest range of molecular probes and reagents to perform in vivo as well as in vitro studies. Based on our experience with various murine transplantation models, we developed a heterotopic pancreas transplantation model in mice with the intent to analyze mechanisms underlying severe ischemia reperfusion injury-associated early graft damage. In contrast to previously described techniques using suture techniques, herein we describe a new procedure using a non-suture cuff technique. 
In recent years, we have performed more than 300 pancreas transplantations in mice with an overall success rate of &gt;90%, a success rate never described before in mouse pancreas transplantation. The backbone of this non-suture cuff technique for graft revascularization consists of two major steps: (I) pulling the recipient vessel over a polyethylene/polyamide cuff and fixing it with a circumferential ligature, and (II) placing the donor vessel over the everted recipient vessel and fixing it with a second circumferential ligature. The resultant continuity of the endothelial layer results in less thrombogenic lesions with high patency rates and, finally, high success rates.
In this model, arterial anastomosis is achieved by pulling the abdominal aorta of the donor graft over the everted common carotid artery of the recipient animal. Venous drainage of the graft is achieved by pulling the portal vein of the graft over the everted external jugular vein of the recipient. This manuscript provides details and crucial steps of the organ recovery and organ implantation procedures, which will allow researchers with microsurgical skills to perform the transplantation successfully in their laboratories.

Among abdominal solid organ transplantation, pancreatic grafts are prone to develop severe ischemia reperfusion injury-associated graft damage, leading eventually to early graft loss. This protocol describes a model of murine pancreas transplantation using a non-suture cuff technique, ideally suited for analyzing these early, deleterious damages.

Mouse models have several advantages in transplantation research, including easy handling, a variety of genetically well-defined strains, and the ...

A Mouse Model of Vascularized Heterotopic Spleen Transplantation for Studying Spleen Cell Biology and Transplant Immunity

The spleen is a unique lymphoid organ that plays a critical role in the homeostasis of the immune and hematopoietic systems. Patients that have undergone splenectomy regardless of precipitating causes are prone to develop an overwhelming post-splenectomy infection and experience increased risks of deep venous thrombosis and malignancies. Recently, epidemiological studies indicated that splenectomy might be associated with the occurrence of cardiovascular diseases, suggesting that physiological functions of the spleen have not yet been fully recognized. Here, we introduce a mouse model of vascularized heterotopic spleen transplantation, which not only can be utilized to study the function and behavioral activity of splenic immune cell subsets in different biologic processes, but also can be a powerful tool to test the therapeutic potential of spleen transplantation in certain diseases. The main surgical steps of this model include donor spleen harvest, the removal of recipient native spleen, and spleen graft revascularization. Using congenic mouse strains (e.g., mice with CD45.1/CD45.2 backgrounds), we observed that after syngeneic transplantation, both donor-derived splenic lymphocytes and myeloid cells migrated out of the graft as early as post-operative day 1, concomitant with the influx of multiple types of recipient cells, thus generating a unique chimera. &#160;Despite relatively challenging techniques, this procedure can be performed with &#62;90% success rate. This model allows tracking the fate, longevity, and function of splenocytes during steady state and in a disease setting following a spleen transplantation, thereby offering a great opportunity to discover the distinct role for spleen-derived immune cells in different disease processes.

This protocol details the surgical steps of a mouse model of vascularized heterotopic spleen transplantation, a technically challenging model that can serve as a powerful tool in studying the fate and longevity of spleen cells, the mechanisms of distinct spleen cell populations in disease progression, and transplant immunity.

The spleen is a unique lymphoid organ that plays a critical role in the homeostasis of the immune and hematopoietic systems. Patients that have undergone ...

A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of transplant with the potential to deliver durable heart valves capable of somatic growth with no requirement for anticoagulation. However, the immunobiology of heart valve transplants remains unexplored, highlighting the need for animal models to study this new type of transplant. Previous rat models for heterotopic aortic valve transplantation into the abdominal aorta have been described, though they are technically challenging and costly. For addressing this challenge, a renal subcapsular transplant model was developed in rodents as a practical and more straightforward method for studying heart valve transplant immunobiology. In this model, a single aortic valve leaflet is harvested and inserted into the renal subcapsular space. The kidney is easily accessible, and the transplanted tissue is securely contained in a subcapsular space that is well vascularized and can accommodate a variety of tissue sizes. Furthermore, because a single rat can provide three donor aortic leaflets and a single kidney can provide multiple sites for transplanted tissue, fewer rats are required for a given study. Here, the transplantation technique is described, providing a significant step forward in studying the transplant immunology of heart valve transplantation.

This protocol describes a simple and efficient method for the transplantation of aortic valve leaflets under the renal capsule to allow for the study of alloreactivity of heart valves.

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of ...

A Porcine Heterotopic Heart Transplantation Protocol for Delivery of Therapeutics to a Cardiac Allograft

Cardiac transplantation is the gold standard treatment for end-stage heart failure. However, it remains limited by the number of available donor hearts and complications such as primary graft dysfunction and graft rejection. The recent clinical use of an ex vivo perfusion device in cardiac transplantation introduces a unique opportunity for treating cardiac allografts with therapeutic interventions to improve function and avoid deleterious recipient responses. Establishing a translational, large-animal model for therapeutic delivery to the entire allograft is essential for testing novel therapeutic approaches in cardiac transplantation. The porcine, heterotopic heart transplantation model in the intraabdominal position serves as an excellent model for assessing the effects of novel interventions and the immunopathology of graft rejection. This model additionally offers long-term survival for the pig, given that the graft is not required to maintain the recipient's circulation. The aim of this protocol is to provide a reproducible and robust approach for achieving ex vivo delivery of a therapeutic to the entire cardiac allograft prior to transplantation and provide technical details to perform a survival heterotopic transplant of the ex vivo perfused heart.

We present a protocol for utilizing a normothermic ex vivo sanguinous perfusion system for the delivery of therapeutics to an entire cardiac allograft in a porcine heterotopic heart transplant model.

Cardiac transplantation is the gold standard treatment for end-stage heart failure. However, it remains limited by the number of available donor hearts ...