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>

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 border="0">
	<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 500ml solution add the ingredient listed below to a 330ml of double distilled water. Mix well, and then fill in the rest of the 170ml of double distilled water into the solution to a final volume of 500ml. 
			Ingredients: 1.02g KH2PO4, 3.66g K2HPO4, 0.56g KCl, 0.42g NaHCO3, and 17.52g of glucose.</td>
		</tr>
		<tr>
			<td>Suture</td>
			<td>Ethilon</td>
			<td>MWI 72667</td>
			<td>6-0 Ethilon</td>
		</tr>
		<tr>
			<td>Suture</td>
			<td>Microsurgery Instruments, Inc.</td>
			<td>10-0 Nylon</td>
			<td>10-0 Nylon</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&#34;&#160;</td>
		</tr>
		<tr>
			<td>Microsurgical dilator</td>
			<td>S&#38;T</td>
			<td>D-5a.1</td>
			<td>Dilator, 11cm, FH, 0.1mm AT10d</td>
		</tr>
	</tbody>
</table>

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...

Watch this Scientific Journal Video about A Novel Microsurgical Model for Heterotopic, En Bloc Chest Wall, Thymus, and Heart Transplantation in Mice at JoVE.com

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

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 ...

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9.2K Views.  Johns Hopkins University School of Medicine. 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.

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

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 ...

Rat Heterotopic Abdominal Heart/Single-lung Transplantation in a Volume-loaded Configuration

Herein, we describe a novel technique for heterotopic abdominal heart-lung transplantation (HAHLT) in rats. The configuration of the transplant graft involves anastomosis of donor inferior vena cava (IVC) to recipient IVC, and donor ascending aorta (Ao) to recipient abdominal Ao. The right upper and middle lung lobes are preserved and function as conduits for blood flow from right heart to left heart.
There are several advantages to using this technique, and it lends itself to a broad range of applications. Because the graft is transplanted in a configuration that allows for dyamic volume-loading, cardiac function may be directly assessed in vivo. The use of pressure-volume conductance catheters permits characterization of load-dependent and load-independent hemodynamic parameters. The graft may be converted to a loaded configuration by applying a clamp to the recipient&#8217;s infra-hepatic IVC. We describe modified surgical techniques for both donor and recipient operations, and an ideal myocardial protection strategy. Depending on the experimental aim, this model may be adapted for use in both acute and chronic studies of graft function, immunologic status, and variable ventricular loading conditions. The conducting airways to the transplanted lung are preserved, and allow for acute lung re-ventilation. This facilitates analysis of the effects of the mixed venous and arterial blood providing coronary perfusion to the graft.
A limitation of this model is its technical complexity. There is a significant learning curve for new operators, who should ideally be mentored in the technique. A surgical training background is advantageous for those wishing to apply this model. Despite its complexity, we aim to present the model in a clear and easily applicable format. Because of the physiologic similarity of this model to orthotopic transplantation, and its broad range of study applications, the effort invested in learning the technique is likely to be worthwhile.

We describe a novel technique for heterotopic abdominal heart-lung transplantation (HAHLT) in rats. The transplant configuration results in a partially loaded graft circulation, allowing direct functional assessment. This model may be employed for acute or chronic studies of function and immunologic status of the transplanted graft.

Herein, we describe a novel technique for heterotopic abdominal heart-lung transplantation (HAHLT) in rats. The configuration of the transplant graft ...

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 ...

Heterotopic Renal Autotransplantation in a Porcine Model: A Step-by-Step Protocol

Kidney transplantation is the treatment of choice for patients suffering from end-stage renal disease. It offers better life expectancy and higher quality of life when compared to dialysis. Although the last few decades have seen major improvements in patient outcomes following kidney transplantation, the increasing shortage of available organs represents a severe problem worldwide. To expand the donor pool, marginal kidney grafts recovered from extended criteria donors (ECD) or donated after circulatory death (DCD) are now accepted for transplantation. To further improve the postoperative outcome of these marginal grafts, research must focus on new therapeutic approaches such as alternative preservation techniques, immunomodulation, gene transfer, and stem cell administration.
Experimental studies in animal models are the final step before newly developed techniques can be translated into clinical practice. Porcine kidney transplantation is an excellent model of human transplantation and allows investigation of novel approaches. The major advantage of the porcine model is its anatomical and physiological similarity to the human body, which facilitates the rapid translation of new findings to clinical trials. This article offers a surgical step-by-step protocol for an autotransplantation model and highlights key factors to ensure experimental success. Adequate pre- and postoperative housing, attentive anesthesia, and consistent surgical techniques result in favorable postoperative outcomes. Resection of the contralateral native kidney provides the opportunity to assess post-transplant graft function. The placement of venous and urinary catheters and the use of metabolic cages allow further detailed evaluation. For long-term follow-up studies and investigation of alternative graft preservation techniques, autotransplantation models are superior to allotransplantation models, as they avoid the confounding bias posed by rejection and immunosuppressive medication.

Porcine models of organ transplantation provide an important platform to study mechanisms of organ preservation. This article describes a heterotopic porcine renal autotransplantation model, which allows investigating new approaches to improve the outcome of transplantation using marginal kidney grafts.

Kidney transplantation is the treatment of choice for patients suffering from end-stage renal disease. It offers better life expectancy and higher quality ...

Cardiopulmonary Bypass in a Mouse Model: A Novel Approach

As prolonged cardiopulmonary bypass becomes more essential during cardiac interventions, an increasing clinical demand arises for procedure optimization and for minimizing organ damage resulting from prolonged extracorporal circulation. The goal of this paper was to demonstrate a fully functional and clinically relevant model of cardiopulmonary bypass in a mouse. We report on the device design, perfusion circuit optimization, and microsurgical techniques. This model is an acute model, which is not compatible with survival due to the need for multiple blood drawings. Because of the range of tools available for mice (e.g., markers, knockouts, etc.), this model will facilitate investigation into the molecular mechanisms of organ damage and the effect of cardiopulmonary bypass in relation to other comorbidities.

This paper describes how to perform cardiopulmonary bypass in mice. This novel model will facilitate the investigation of the molecular mechanisms involved in organ damage.

As prolonged cardiopulmonary bypass becomes more essential during cardiac interventions, an increasing clinical demand arises for procedure optimization ...

A Pre-Clinical Porcine Model of Orthotopic Heart Transplantation

Fifty-years following the first successful report, cardiac transplantation remains the gold-standard treatment for eligible patients with advanced heart failure. Multiple small-animal models of heart transplantation have been used to study the acute and long-term effects of novel therapies. However, few are tested and demonstrated success in clinical trials. It is of critical importance to evaluate new therapies in a clinically relevant large-animal model for efficient and reliable translation of basic studies&#39;&#160;findings. Here, we describe a pre-clinical large-animal (porcine) model of orthotopic heart transplantation that has been firmly established and previously used to investigate novel cardioprotective strategies. This procedure focuses on acute ischemia-reperfusion injury and is a reliable method to investigate novel interventions which have been tested and validated in smaller experimental models, such as the murine model. We demonstrate its usefulness in assessing cardiac performance during the early post-transplantation period and other potential possibilities enabled by the model.

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.

Fifty-years following the first successful report, cardiac transplantation remains the gold-standard treatment for eligible patients with advanced heart ...

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 ...