Name: A Novel Microsurgical Model for Heterotopic, En Bloc Chest Wall, Thymus, and Heart Transplantation in Mice (Video) | JoVE Journal
Uploaded: 2016-01-23T14:00:00.000+00:00
Duration: 12 min 24 s
Description: Learn a novel murine model for heterotopic chest wall, thymus, and heart transplantation using cuff anastomosis techniques. This method enables researchers to investigate immune tolerance mechanisms and improve allograft survival in reconstructive transplantation.

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5 minutes

This work was funded by the American Association of Plastic Surgeons 2014 Academic Scholar Award.

All operative procedures were completed in compliance with Johns Hopkins University and the United States Department of Agriculture and Public Health Service requirements. This protocol follows the Johns Hopkins University Animal Care and Use Committee, institutional review board approved guidelines (protocol number M013M490). Final survival data was recorded for the surgical procedures described below.&#160;Both donor and recipient animals receive pre-emptive anesthesia using buprenorphine at 0.1 mg/kg s.c. one hour prior to surgery and in the recipient animal buprenorphine is re-administered at the same dose after transplant and re-dosed as needed in the first 48 hours after surgery.&#160;
1. Donor Allograft Recovery
Note: Begin the donor portion of the transplant 40 min&#160;earlier than the recipient transplant to minimize recipient anesthesia time and to facilitate a simultaneous end time or slightly earlier end time versus the recipient preparation.
<ol>
	<li>Use standard sterile microsurgical instruments and sterile gloves for the procedure. Our laboratory uses autoclave sterilization of microsurgical instruments.</li>
	<li>Anesthetize the donor mouse (male) using isoflurane induction vaporizer at 4%. Using atraumatic mechanical clippers remove the hair from the cervical, thoracic, and abdominal region. Place the animal in the supine position and maintain isoflurane on 1-2% through a nose cone. Ensure adequate anesthesia throughout the procedure by periodically evaluating the toe pinch withdrawal reflex.</li>
	<li>Prior to the skin incision, widely prepare the operative by applying povidone iodine antiseptic followed by isopropyl alcohol using a sterile cotton swab.</li>
	<li>Begin with a superficial transverse skin incision with scissors across the cervical and abdominal skin. Connect both incisions bilaterally along the midaxillary lines.</li>
	<li>Using microsurgical forceps dissect the cervical region bilaterally to identify, ligate and divide the external jugular veins with 6-0 silk suture and scissors. Then using electrocautery divide the sternocleidomastoid muscles to expose the internal jugular veins and common carotid arteries, bilaterally. Pass a 6-0 silk suture under the left-sided and the right-sided common carotid and internal jugular veins in bulk fashion. 
	NOTE: They will be tied and divided later in Step 1.9.</li>
	<li>Sharply divide the strap muscles and associated loose areolar tissue, located anterior to the trachea, using scissors to free the remaining attachments of the cervical region.</li>
	<li>Using bipolar electrocautery and sharp dissection, divide the pectoralis major muscles and clavicles to expose the subclavian vessels and ligate (6-0 silk suture) and divide proximally.</li>
	<li>Next, gently, grasp and withdraw the animal&#8217;s penis. Along the dorsum of the penis visualize the dorsal vein of the penis, and disinfect the region with isopropyl alcohol. Using a 30 G needle, inject 30,000 units of heparin intravenously through the dorsal vein and allow the penis to recoil back to its original position.&#160;&#160;Partial leakage of the heparin solution into the surrounding tissue may occur.&#160;</li>
	<li>Using the previously placed bulk ties around common carotid artery and internal jugular vein, ligate and divide the structures, bilaterally.</li>
	<li>Next use scissors to make a transverse intrabdominal incision. Eviscerate the intestines to expose the infrahepatic inferior vena cava and inject 2 ml of cold Euro-Collins cardioplegia solution into the infrahepatic inferior vena cava. Ensure proper injection by visualizing liver discoloration and cessation of the heartbeat prior to advancing to the following step. 
	NOTE: Euro-Collins solution is prepared in our laboratory, see table of specific reagents and instruments.</li>
	<li>Using scissors access the intrathoracic cavity via a bilateral diaphragmatic incision from the exposed abdomen. Extend the incision cephalad through the intercostals muscles and ribs. Reflect chest wall exposing the heart, thymus, and great vessels while simultaneously ensuring preservation of the internal thoracic vessels along the chest wall.</li>
	<li>Inject the suprahepatic inferior vena cava with 4 ml of cold Euro-Collins cardioplegia solution.</li>
	<li>Identify the root of the aorta and trace distally to the descending aorta. Sharply cut the descending aorta (preserving maximal length).</li>
	<li>Identify the pulmonary trunk and divide just proximal to its branch point (preserving maximal length). Then using 2 ml of cold Euro-Collins cardioplegia solution, flush the pulmonary trunk and heart by placing a soft plastic tip catheter into the lumen of the pulmonary trunk.</li>
	<li>Using a 6-0 silk suture, ligate and divide the inferior vena cava, confluence of pulmonary veins, and accessory branches of the bilateral superior vena cava. Then elevate and dissect the heart cephalad from the attachments along the main stem bronchi and trachea with care not to enter the airway. Using sharp and bipolar electrocautery dissect the chest wall, thymus, and heart completely liberating it from the donor mouse.</li>
	<li>Finally, trim the allograft chest wall ex vivo to a smaller size, using scissors, along the sternum and lateral costae, with care not to disrupt the internal thoracic vessels (Figure 1A). To minimize hemorrhage following revascularization, use bipolar electrocauterization along the borders of the osteomusculocutaneous sternum.</li>
	<li>Place the allograft in 10 ml of cold (4o Celcius) Euro-Collins solution if the recipient is not prepared for inset. However, if the recipient is ready for inset, transfer the allograft straight to the recipient operative field.</li>
</ol>
2. Recipient Preparation
Note: To minimize recipient anesthesia time, begin the recipient preparation at a separate operative station approximately 40 min&#160;prior to completion of the donor allograft harvest.
<ol>
	<li>Use a separate set of standard sterile microsurgical instruments and sterile gloves for the procedure.</li>
	<li>Anesthetize the recipient mouse (male or female) using isoflurane induction vaporizer at 4%. Using atraumatic mechanical clippers remove the hair from the right cervical and thoracic region.</li>
	<li>Place the mouse in the supine position and angle the right upper limb slightly inferiorly forming a 110 degree angle between the head and right upper limb. Maintain anesthesia on 1-2% isoflurane through a nose cone.</li>
	<li>Place petroleum ophthalmic ointment on the mouse eyes using a cotton tip applicator. Prior to the skin incision, widely prepare the operative site using povidone iodine antiseptic followed by isopropyl alcohol.</li>
	<li>Using scissors, make a skin incision from the midline along the right inferior border of the mandible and extend the incision infero-laterally to the right thoracic region. Using blunt dissection with microvascular forceps, mobilize the external jugular vein by circumferentially free the vessel from soft tissue and adventitia. Divide all branches using electrocautery, and remove the right lobe of the submandibular gland using sharp dissection and electrocautery to free space for the allograft.</li>
	<li>Ensure enough length of the external jugular vein to evert over a cuff, and ligate the external jugular vein using a 6-0 silk suture. Insert the vein through the lumen of a precut polyimide cuff and use a bulldog microvascular clamp to fix the vessel-cuff complex in place. Then using scissors, proximally divide the external jugular vein, evert over the cuff, and fix in place with a 10-0 nylon suture. (Figure 1B)</li>
	<li>Divide the right sternocleidomastoid muscle with bipolar electrocautery to expose the common carotid artery. Circumferentially mobilize the artery cephalad to the distal most point within the cervical region. This is accomplished using blunt dissection of the vessel with forceps to remove soft tissue and surrounding adventitia.</li>
	<li>Using 6-0 silk suture, ligate and divide the common carotid artery. Pass the artery through the lumen of a precut polyimide cuff and fix it in place with a bulldog microvascular clamp as close to the thoracic inlet as possible. Divide the vessel distally, gently dilate the vessel using a microsurgical dilator, evert over the cuff, and fix in place with a 10-0 nylon suture. (Figure 1B) 
	NOTE: The specific microsurgical dilator is described in the table of specific reagents and instruments.</li>
</ol>
3. Allograft Inset
<ol>
	<li>Maintain standard sterile instrumentation and sterile gloves to place the allograft within the recipient cervical region in an upside down and oblique position.</li>
	<li>Next, place the donor descending aortic lumen over the arterial cuff construct of the recipient and fix it in place it with a 10-0 nylon suture (Figure 1C and 1D).</li>
	<li>Fashion the same anastomosis as in step 3.2 between the donor pulmonary artery and the everted external jugular vein-cuff construct of the recipient mouse (Figure 1C and 1D).</li>
	<li>First remove venous microvascular clamp (external jugular vein clamp) and then release the arterial clamp (common carotid artery clamp). During arterial reperfusion, inspect the entirety of the allograft to address any hemorrhage. If hemorrhage is visualized, reapply the arterial clamp to minimize blood loss and mitigate the source of bleeding using bipolar electrocautery.</li>
	<li>Inspect the graft and ensure hemostasis. Release and completely remove the arterial microvascular clamp. Observe the heart to show signs of reperfusion, which will be instantaneously apparent with rapid volume expansion of the heart chambers, and wait for beating to begin within 0.5-1 min. Use warm saline (35&#176; Celsius) to moisten the heart.</li>
	<li>Drape the chest wall into an anatomical position so as not to induce any kinking or tensions on the anastomoses. Close the skin of the surgical wound is using 6-0 continuous nylon sutures (Figure 1E).</li>
</ol>
4. Postoperative Care
<ol>
	<li>Administer a 0.3 ml normal saline intraperitoneal fluid bolus immediately postoperatively for fluid replacement.</li>
	<li>Then subcutaneously inject buprenorphine (0.1mg/kg) and enrofloxacin (5mg/kg) for pain and infection prophylaxis, respectively.</li>
	<li>Place the animal under a heat lamp until awaking from anesthesia and return to sternal recumbency. During recovery, inspect the neck to visualize the fibrillating heartbeat of the allograft ensuring adequate allograft perfusion.</li>
	<li>Once awake and in the recumbent position, return the mouse to a separate cage (without the company of other mice) where it can receive food and water ad libitum. Due to any temporary minor restrictive motion of the right upper limb, leave a gelatin food source on the floor of the cage.</li>
	<li>Observe the recipient mouse for 1 hr postoperatively and then return it to the cage facility where it can receive food and water ad libitum and is inspected three times a day for the first 24 hr for activity and nutritional intake. Monitor mice for signs of pain and distress and re-dose with buprenorphine (0.1mg/kg) subcutaneously twice a day as needed for the first 72 hr. Examine the animals daily thereafter and weigh them every week.</li>
	<li>Consult with a veterinary staff member if any mice display signs of pain, distress, or decreased feed intake. Consider early euthanasia (in our protocol the euthanasia technique employs CO2 overdose for 7 min, followed by cervical dislocation).</li>
	<li>Cessation of the allograft heart beat is defined as a specific endpoint prompting the mouse to be sacrificed.</li>
</ol>

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<li>Leventhal, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Chimerism+and+tolerance+without%2C+GVHD+or+engraftment+syndrome+in%2C+HLA-mismatched+combined+kidney+and+hematopoietic+stem+cell+transplantation%5BTitle%5D)+AND+%22Sci+Transl+Med%22%5BJournal%5D)">Chimerism and tolerance without, GVHD or engraftment syndrome in, HLA-mismatched combined kidney and hematopoietic stem cell transplantation.</a> Sci Transl Med. 4 (124), 124-128 (2012).</li>
<li>Talmor, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Bone+marrow-derived+chimerism+in+non-irradiated%2C+cyclosporin-treated+rats+receiving+microvascularized+limb+transplants%3A+evidence+for+donor-derived+dendritic+cells+in+recipient+lymphoid+tissues%5BTitle%5D)+AND+%22Immunology%22%5BJournal%5D)">Bone marrow-derived chimerism in non-irradiated, cyclosporin-treated rats receiving microvascularized limb transplants: evidence for donor-derived dendritic cells in recipient lymphoid tissues.</a> Immunology. 86 (3), 448-455 (1995).</li>
<li>Arslan, E., Klimczak, A., Siemionow, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Chimerism+induction+in+vascularized+bone+marrow+transplants+augmented+with+bone+marrow+cells%5BTitle%5D)+AND+%22Microsurgery%22%5BJournal%5D)">Chimerism induction in vascularized bone marrow transplants augmented with bone marrow cells.</a> Microsurgery. 27 (3), 190-199 (2007).</li>
<li>Page, E. K., Dar, W. A., Knechtle, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Tolerogenic+therapies+in+transplantation%5BTitle%5D)+AND+%22Front+Immunol%22%5BJournal%5D)">Tolerogenic therapies in transplantation.</a> Front Immunol. 3, 198(2012).</li>
<li>Lukomska, B., Durlick, M., Cybulska, E., Olszewski, W. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Comparative+analysis+of+immunological+reconstitution+induced+by+vascularized+bone+marrow+versus+bone+marrow+cell+transplantation%5BTitle%5D)+AND+%22Transpl+Int%22%5BJournal%5D)">Comparative analysis of immunological reconstitution induced by vascularized bone marrow versus bone marrow cell transplantation.</a> Transpl Int. 9, Suppl 1. S492-S496 (1996).</li>
<li>Brandacher, G., Lee, W. P., Schneeberger, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Minimizing+immunosuppression+in+hand+transplantation%5BTitle%5D)+AND+%22Expert+Rev+Clin+Immunol%22%5BJournal%5D)">Minimizing immunosuppression in hand transplantation.</a> Expert Rev Clin Immunol. 8 (7), 673-683 (2012).</li>
<li>Lin, C. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+neck+as+a+preferred+recipient+site+for+vascularized+composite+allotransplantation+in+the+mouse%5BTitle%5D)+AND+%22Plast+Reconstr+Surg%22%5BJournal%5D)">The neck as a preferred recipient site for vascularized composite allotransplantation in the mouse.</a> Plast Reconstr Surg. 133 (2), 133e(2014).</li>
<li>Sucher, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Hemiface+allotransplantation+in+the+mouse%5BTitle%5D)+AND+%22Plast+Reconstr+Surg%22%5BJournal%5D)">Hemiface allotransplantation in the mouse.</a> Plast Reconstr Surg. 129 (4), 867-870 (2012).</li>
<li>Sucher, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mouse+hind+limb+transplantation%3A+a+new+composite+tissue+allotransplantation+model+using+nonsuture+supermicrosurgery%5BTitle%5D)+AND+%22Transplantation%22%5BJournal%5D)">Mouse hind limb transplantation: a new composite tissue allotransplantation model using nonsuture supermicrosurgery.</a> Transplantation. 90 (12), 1374-1380 (2010).</li>
<li>Sucher, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Orthotopic+hind-limb+transplantation+in+rats%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Orthotopic hind-limb transplantation in rats.</a> J Vis Exp. (41), e2022(2010).</li>
<li>Maglione, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+novel+technique+for+heterotopic+vascularized+pancreas+transplantation+in+mice+to+assess+ischemia+reperfusion+injury+and+graft+pancreatitis%5BTitle%5D)+AND+%22Surgery%22%5BJournal%5D)">A novel technique for heterotopic vascularized pancreas transplantation in mice to assess ischemia reperfusion injury and graft pancreatitis.</a> Surgery. 141 (5), 682-689 (2007).</li>
<li>Zou, Y., Brandacher, G., Margreiter, R., Steurer, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cervical+heterotopic+arterialized+liver+transplantation+in+the+mouse%5BTitle%5D)+AND+%22J+Surg+Res%22%5BJournal%5D)">Cervical heterotopic arterialized liver transplantation in the mouse.</a> J Surg Res. 93 (1), 97-100 (2000).</li>
<li>Kamada, N., Calne, R. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Orthotopic+liver+transplantation+in+the+rat.+Technique+using+cuff+for+portal+vein+anastomosis+and+biliary+drainage%5BTitle%5D)+AND+%22Transplantation%22%5BJournal%5D)">Orthotopic liver transplantation in the rat. Technique using cuff for portal vein anastomosis and biliary drainage.</a> Transplantation. 28 (1), 47-50 (1979).</li>
<li>Zimmermann, F. A., Butcher, G. W., Davies, H. S., Brons, G., Kamada, N., Türel, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Techniques+for+orthotopic+liver+transplantation+in+the+rat+and+some+studies+of+the+immunologic+responses+to+fully+allogeneic+liver+grafts.+Transplant%5BTitle%5D)+AND+%22Proc%22%5BJournal%5D)">Techniques for orthotopic liver transplantation in the rat and some studies of the immunologic responses to fully allogeneic liver grafts. Transplant.</a> Proc. 11 (1), 571-577 (1979).</li>
</ol>

The authors do not have any conflicts of interest or financial disclosures to declare.

There are a multitude of phenomena that factor into the immunological investigation of allotransplantation, which include but are not limited to mechanisms of acute and chronic rejection, direct and indirect antigen presentation, recipient sensitization, or the induction of mixed chimerism.19 Animal models have become the gold standard for the study of transplant immunology, and mouse models are popularly implemented due to their low cost, availability of transgenic and gene knockout mice, commercially availab...

Cardiac transplantation is the treatment of choice for end-stage heart failure. Both technical advancements and pharmacological innovations have propelled the field to early graft acceptance rates above 90%.12,13 Despite this, 60-80% 5-year graft survival is at a standstill and chronic rejection, characterized by transplant vasculopathy, remains inevitable.14-16 Furthermore, patients are subjected to multiple surgical procedures and lifelong immunosuppression, which are associated with chest wall deformities and medical sequelae and toxicities, respectively. The need for innovative approaches to extend allograft survival, minimize the immunosuppressive requirements, and offer reconstructive options for anatomical deformities is pressing.
Vascularized composite allotransplantation offers a unique strategy for improving heart transplant outcomes both from an immunological aspect as well as a reconstructive perspective.17 Vascularized composite allografts are also unique in a way that they have an inherent source of donor-derived hematopoietic stem cells which has shown a favorable ability to reduce immunosuppression and induce and sustain mixed chimerism.1-8 Additionally, co-transplantation of the thymus has shown to prolong survival of both, solid organ transplants and vascularized composite allografts.2,9-11 Combining these strategies with heart transplantation offers a novel solution to the aforementioned challenges facing heart transplantation.18
Murine models serve as excellent platforms for mechanistic in vivo investigation because of the availability of antibodies and well-defined inbred and knockout strains.19-21 Although heart transplantation in mice is commonly studied using a heterotopic intraabdominal microsurgical suture transplant model22-25, a heterotopic, cervical, non-suture cuff technique model has shown to be extremely replicable, reliable, and carries fewer rates of thrombosis.19,26,27 The goal of this study is to develop a heterotopic en bloc osteomyocutaneous chest wall, thymus, and heart transplant technique in mice to study the immunological mechanisms of combined solid organ and vascularized composite allotransplantation using a cervical non-suture cuff technique. This cluster allograft is perfused through the anastomosis of the donor descending aorta to the right common carotid artery and the donor pulmonary artery to the right external jugular vein. Preservation of the internal thoracic vessels and associated thymus branches is paramount to perfusing the chest wall (sternum, ribs, muscles, and skin) and thymus.

<table><tbody><tr><td>Euro-Collins Solution</td><td></td><td></td><td>The solution is not commercially purchased but rather prepared in the laboratory. To make a 500 ml solution add the ingredient listed below to a 330 ml of double distilled water. Mix well, and then fill in the rest of the 170 ml of double distilled water into the solution to a final volume of 500 ml.&lt;br/&gt; Ingredients: 1.02 g KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, 3.66 g K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, 0.56 g KCl, 0.42 g NaHCO&lt;sub&gt;3&lt;/sub&gt;, and 17.52 g of glucose.</td></tr><tr><td>Suture</td><td>Ethilon</td><td>MWI 72667</td><td>6-0 Ethilon


https://www.mwivet.com (MWI - Veterinary Supplies)</td></tr><tr><td>Polyimide Cuff Vein (21G)</td><td>Vention Medical</td><td>141-0043</td><td>http://www.ventionmedical.com/products-and-services/polyimide-tubing/</td></tr><tr><td>Polyimide Cuff Artery (24G)</td><td>Vention Medical</td><td>141-0027</td><td>http://www.ventionmedical.com/products-and-services/polyimide-tubing/</td></tr><tr><td>Soft plastic tip catheter</td><td>Terumo</td><td>SR*OX2419CA</td><td>24G x 3/4&quot;&amp;nbsp;</td></tr><tr><td>Microsurgical dilator</td><td>S&amp;amp;T</td><td>D-5a.1</td><td>Dilator, 11 cm, FH, 0.1 mm AT10d</td></tr></tbody></table>

a novel microsurgical model for heterotopic, en bloc chest wall, thymus, and heart transplantation in mice

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.

Syngeneic C57BL/6 transplants achieved long term survival. The design of the allograft (Figure 1) proved to be successful from an animal survival perspective and the ability to evaluate ongoing allograft survival. This was demonstrated through overlying skin remaining viable, active ongoing allograft hair growth, and heartbeats were able to be evaluated with visualization and finger palpation. Survival data is represented in Figure 2 for syngeneic transplanted mice. The mean survival tim...

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 ...

The overall goal of this protocol is to introduce a novel murine model of solid organ and vascularized composite allotransplantation to investigate the effects of donor-derived vascularized hematopoietic and primary lymphoid tissue on immune tolerance and prolonged allograft survival. This method can help answer key questions in the field of reconstructive transplantion such as the role of vascularized bone and thymus on tolerance induction. The main advantage of this technique is that for the first time, combined vascularized bone marrow and thymus is being transplanted with a solid organ, the heart and the immunologically versatile mass model system.The implications of this technique extend from the basic immunological research toward therapeutic translation into the clinical arena to treat complex tissue defects and simultaneously improve long-term survival in solid organ transplantion. Demonstrating this procedure will be Dr.BC Oh and Dr.Georg Furtmuller, who are post-doctoral fellows in the department of plastic and reconstructive surgery at the Johns Hopkins University. To begin, after anesthetizing a male donor mouse according to the text protocol, while wearing sterile gloves, use mechanical clippers to remove the hair from the cervical, thoracic and abdominal region.On a sterile surgical surface, place the animal in a supine position. Attach a nose cone and maintain isoflurane at 1 to 2%Prior to the skin incision, widely prepare the incision site by using a sterile cotton swab to apply povidone iodine antiseptic followed by isopropyl alcohol. Using scissors, make superficial transfer of skin incisions across the cervical and abdominal skin.Connect both incisions bilaterally, along the midaxillary lines. With microforceps, dissect the cervical region bilaterally to identify the external jugular veins. And with 6-0 silk suture, ligate the vessels before using scissors to divide them.Then with electrocautery, divide the sternocleidomastoid muscles to bilaterally expose the internal jugular veins and common carotid arteries. Pass a 6 0 silk suture under the left and right common carotid arteries and internal jugular veins, respectively, in bulk fashion. Using bipolar electrocautery and sharp dissection, divide the pectoralis major muscles and clavicles to expose the subclavian vessels.Used 6-0 silk suture to ligate and divide the vessels proximally. Next, gently grasp and withdraw the animal's penis. Along the dorsum of the penis, locate the dorsal vein and use isopropyl alcohol to disinfect the region.Then, using a 30-gauge needle, inject 30, 000 units of heparin intravenously through the dorsal vein and allow the penis to recoil back to its original position. With the previously placed bulk ties around the common carotid artery and internal jugular vein, ligate and divide the structures bilaterally. Now use scissors to make a transverse incision to access the abdominal cavity.Eviscerate the intestines to expose the intrahepatic inferior vena cava and inject two milliliters of cold EuroCollins cardioplegia solution into the intrahepatic inferior vena cava. Ensure proper injection by visualizing liver discoloration and cessation of the heartbeat. Using scissors, access the intrathoracic cavity via a bilateral diaphragmatic incision from the exposed abdomen, extending the incision cephalad through the intercostal muscles and ribs.Flossing the pulmonary trunk and heart by using EuroCollin solution is critical to prevent micro thromboembolism from potential remaining inside of the coronary circulation. After injecting cold EuroCollins solution into the super hepatic inferior vena cava according to the text protocol, identify the root of the aorta and trace distally to the descending aorta. Sharply cut the descending aorta, preserving maximal length.Now identify the pulmonary trunk and divide just proximal to its branch point, preserving maximal length. Then using two milliliters of cold EuroCollins cardioplegia solution, flush the pulmonary trunk and heart by placing a soft plastic tape catheter into the lumen of the pulmonary trunk. Using 6-0 silk suture, ligate and divide the inferior vena cava, confluence of pulmonary veins and accessory branches of the bilateral superior vena cava.Then evaluate and dissect the heart cephalad from the attachments along the main stem bronchi and trachea with care, so as not to enter the airway. With sharp and bipolar electrocautery, dissect the chest wall, thymus and heart, completely liberating it from the donor mouse. Finally, using scissors, trim the allograft chest wall ex vivo to a smaller size along the sternum and lateral costi, taking care not to disrupt the internal thoracic vessels.To minimize hemorrhage following revascularization, use bipolar electrocauterization along the borders of the osteomusculocutaneous sternum. After preparing the recipient mouse according to the text protocol, place the mouse in a supine position and angle the right upper limb with a slightly inferior orientation, forming a 110 degree angle between the head and the right upper limb. Make an incision from the midline along the right inferior border of the mandible, and extend the incision infralaterally to the right thoracic region.With blunt dissection using microvascular forceps, mobilize the external jugular vein by circumferentially freeing the vessel from the soft tissue and adventitia. Use electrocautery to divide all branches and remove the right lobe of the submandibular gland with sharp dissection and electrocautery to free the space for the allograft. Ensure enough length of the jugular vein to evert over a cuff and ligate the external jugular vein using 6-0 silk suture.Then insert the vein through the lumen of a precut polyamide cuff and use a bulldog microvascular clamp to fix the vessel cuff complex in place. Evert over the cuff and use 10-0 nylon suture to fix in place. While carrying out bipolar electrocautery, divide the right sternocleidomastoid muscle to expose the common carotid artery.With forceps and blunt dissection, circumferentially mobilize the arteries cephalad to the distal most point within the cervical region by removing soft tissue and surrounding adventitia. Using 6-0 silk suture, ligate and divide the common carotid artery. Pass the artery through the lumen of a precut polyamide cuff and use a bulldog microvascular clamp to fix it in place as close to the thoracic inlet as possible.Then divide the vessel distally. Use a microsurgical dilator to gently dilate the vessel, evert over the cuff and use 10-0 nylon suture to fix in place. While maintaining sterile conditions, place the donor descending aortic lumen over the arterial cuff construct on the recipient and use 10-0 nylon suture to fix it in place.Fashion a similar anastomosis between the donor pulmonary artery and the everted external jugular vein cuff construct of the recipient mouse. Then remove the Venus microvascular clamp followed by the arterial clamp. Next, inspect the entire allograft for hemorrhage and, if present, mitigate bleeding according to the text protocol.Inspect the graft and ensure hemostasis before releasing and completely removing the arterial microvascular clamp. Observe the heart to identify signs of reperfusion, which will be instantaneously apparent with rapid volume expansion of the heart chambers and wait for beading to begin within 30 to 60 seconds. Use warm saline to moisten the heart then place the chest wall into an anatomical position so as not to induce any kinking or tensions on the anastomoses.Finally, use 6-0 nylon sutures to close the skin of the surgical wound and carry out postoperative care according to the text protocol. Performing end-to-end non-suture cuff anastomosis, maintaining meticulous hemostasis with electrocautery during reperfusion and assuring perfusion of all components of the composite tissue graph are the most critical steps of the procedure. The combined VCA and solid organ allograft is mounted over the previously placed polyamide cuff in the recipient's neck, illustrating optimal graft position and meticulous mounting of the donor vessel onto the cuff's construct.The design of the allograft demonstrated here approved feasible for sustaining perfusion to the chest wall with minimal impact to the recipient animal. Presented here is survival data for syngeneic transplanted mice. The mean survival time was greater than 109 days, suggesting that the technical aspect of the transplanted allograft is designed to perfuse the entirety of the chest wall, thymus and heart.Additionally, this mouse model is not only feasible but can be replicated. Once mastered, this technique can be done in 90 minutes if it is performed in a two-team approach. While attempting this procedure, it's important to remember that hemostasis throughout the entirety of the procedure is paramount, and minimization of kinking during the allograft insetting is also very important.This technique paves the way for researchers in the fields of reconstructive transportation and solid organ transportation to explore the role of vascularized bone marrow and thymus as potential adjuncts to solid organ transportation to improve allograft survival. After watching this video, you should have a good understanding of how to establish a murine model of combined solid organ and vascularized composite allotransportation through a heterotopic unblock chest wall, thymus and heart transplant.

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Research

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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.

Heterotopic Heart Transplantation in Mice

The murine cervical heart transplantation model is well suited for immunological as well as ischemia reperfusion injury studies. We modified the procedure using a non-suture cuff technique and performed more than 1,000 successful transplants with this approach. Herein, we provide additional details of this technique to supplement the video.

Murine Cervical Heart Transplantation Model Using a Modified Cuff Technique

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.

A Modified Method for Heterotopic Mouse Heart Transplantion

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.

Heterotopic Cervical Heart Transplantation in Mice

Here, we describe a pre-clinical large-animal (porcine) model of orthotopic heart transplantation that has been firmly established and utilized to investigate novel cardioprotective strategies.

A Pre-Clinical Porcine Model of Orthotopic Heart Transplantation

A novel technique for blood circuit reconstruction in a heterotopic abdominal mouse heart transplantation model is demonstrated.

Blood Circuit Reconstruction in an Abdominal Mouse Heart Transplantation Model

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.

A Modified Cuff Technique for Mouse Cervical Heterotopic Heart Transplantation Model

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.

Mouse Heterotopic Cervical Cardiac Transplantation Utilizing Vascular Cuffs

This protocol presents a cuff-free, suture-based cervical heterotopic heart transplantation technique in mice, designed to enable the reliable placement of a second heterotopic cardiac graft for evaluating donor-specific immune tolerance under physiological conditions with minimal interference from foreign material.

Cuff-Free Suture-Based Cervical Model for Secondary Heterotopic Heart Transplantation to Evaluate Donor-Specific Immune Tolerance in Mice

This protocol presents a simplified, end-to-side suture anastomosis technique for murine cervical heart transplantation, providing a robust and reproducible model for studying transplant immunology.

Cervical Heterotopic Mouse Heart Transplantation using End-to-side Anastomosis Technique to Study Transplant Immunology

Learn a novel murine model for heterotopic chest wall, thymus, and heart transplantation using cuff anastomosis techniques. This method enables researchers to investigate immune tolerance mechanisms and improve allograft survival in reconstructive transplantation.

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

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