Figure 1 was created with biorender.com. This work was&#160;supported in part&#160;by the AATS Foundation Surgical Investigator Program to TKR, the Children&#39;s Excellence Fund held&#160;by the Department of Pediatrics at the Medical University of South Carolina to TKR, an Emerson Rose Heart Foundation grant to TKR, Philanthropy by Senator Paul Campbell to TKR, NIH-NHLBI Institutional Postdoctoral Training Grants (T32 HL-007260) to JHK and BG, and the Medical University of South Carolina College of Medicine Pre-clerkship FLEX Research Fund to MAH.

The study was approved by the Committee of Animal Research following the National Institutes of Health Guide for Care and Use of Laboratory Animals.
1. Information on the animal model (Rats)
<ol>
	<li>Use an operating microscope (see Table of Materials) with up to 20x magnification for all surgical procedures.</li>
	<li>Use syngeneic (such as Lewis-Lewis) or allogeneic (such as Lewis-Brown Norway) strains for the transplants as needed for the experiment.</li>
	<li>Use rats of age between 5-7 weeks and bodyweight of 100-200 g that are appropriate for the experimental question.</li>
</ol>
2. Removal of fur, preparation of the skin, and anesthesia
<ol>
	<li>Perform all operations under sterile conditions. 
	NOTE: The step is performed in a dedicated surgical space and under sterile conditions.</li>
	<li>Place the rats into an anesthetic induction chamber and induce anesthesia with 5% isoflurane in oxygen. Maintain anesthesia with 3.5% isoflurane in oxygen throughout the procedure.</li>
	<li>For the donor operation, remove the rat&#39;s fur from the umbilicus to the sternal notch using fur clippers. For the recipient operation, clip the hair over the surgical field at the posterior axillary line from the ribs to the pelvis. Next, prepare the skin with a surgical disinfectant.</li>
	<li>Obtain a surgical plane of anesthesia before starting the procedure. Confirm adequate depth of anesthesia by firmly compressing the toes of the rat with forceps. If the rat withdraws to pain, titrate the anesthetic as needed.</li>
	<li>Monitor the respiratory rate and the depth of anesthesia clinically throughout the procedure; the level of isoflurane is adjusted as needed to maintain a breathing rate of 55-65 breaths/min.</li>
</ol>
3. Donor operation
<ol>
	<li>Prepare and anesthetize the rat as stated in step 2. Incise the skin from the xiphoid to the sternal notch using dissecting scissors. Perform a sternectomy by cutting the ribs on each side lateral to the sternum until optimal access to the heart is achieved.</li>
	<li>Heparinize the rat with a 100 U/100 g of injection into the left atrium.</li>
	<li>Sacrifice the donor via exsanguination.</li>
	<li>Excise the thymus to improve the visualization of the great vessels. Then, remove the heart en bloc with the ascending aorta until the level of the innominate artery.</li>
</ol>
4. Preparation of aortic valve leaflets
<ol>
	<li>Place the donor heart in a sterile Petri dish immediately following the cardiectomy. Dissect the donor heart in an ice-cold cold storage buffer (see Table of Materials).</li>
	<li>Using forceps and Vannas spring scissors, dissect the donor heart until only the aortic root remains with a 1 mm ventricular cuff proximal to the aortic valve.</li>
	<li>Open the aortic valve by making a longitudinal cut to open the Sinus of Valsalva between the left and non-coronary sinuses to visualize all three leaflets. 
	NOTE: The cut should be the entire length of the Sinus of Valsalva. The actual dimensions depend on the size of the rat.</li>
	<li>Excise each aortic valve leaflet individually. Specifically, use blunt forceps to grasp the edge of the leaflet and use Vannas spring scissors to excise the leaflet by cutting from one commissure down to the annulus, and then toward the next commissure. 
	NOTE: Take special care to only grasp the edge of the leaflet to minimize disruption of the valvular endothelial cells.</li>
	<li>Store the samples following leaflet excision in ice-cold storage buffer solution until they are ready to be implanted in the recipient rat. Implant all the leaflets within 4 h of cold storage.</li>
</ol>
5. Recipient operation
<ol>
	<li>Prepare and anesthetize the rat as stated in step 2.&#160;Use a heating pad maintained at 36-38 &#176;C to perform the surgery.</li>
	<li>Administer buprenorphine (0.03 mg/kg subcutaneously) to all recipient rats before surgery and every 6-12 h post-operatively as needed to alleviate the pain.</li>
	<li>Place the rat in a right lateral recumbent position to access the left kidney. 
	NOTE: The left kidney is preferred due to its more caudal position relative to the right kidney.</li>
	<li>Incise the skin over the flank longitudinally over 1-inch using scissors. 
	NOTE: The incision must remain smaller than the size of the kidney to provide enough tension to prevent the kidney from retracting back into the abdominal cavity during the procedure.</li>
	<li>Similarly, incise the underlying abdominal wall.</li>
	<li>Externalize the kidney
	<ol>
		<li>Using the thumb and forefinger, apply light pressure dorsally and ventrally while using curved forceps to lift the caudal pole of the kidney through the abdominal and skin incision. Externalize the cranial end of the kidney similarly.</li>
		<li>Alternatively, the kidney may be externalized by grasping the perirenal fat and pulling upward with light tension. 
		NOTE: Take care not to grasp the kidney or the renal vessels directly.</li>
		<li>Once the kidney is externalized, keep it moist with warm saline trickled onto the kidney.</li>
	</ol>
	</li>
	<li>Create a subcapsular pocket.
	<ol>
		<li>Lightly apply pressure to the renal capsule using one set of blunt forceps so that the renal capsule can be clearly distinguished from the underlying parenchyma. Simultaneously using another set of blunt forceps, carefully grasp the capsule and gently pull upward to create a hole in the capsule. 
		NOTE: Due to the delicate nature of the capsule, minimal force is required to establish this incision.</li>
		<li>Continue using blunt forceps to extend the incision until a ~2mm space has been created to accommodate the aortic valve leaflet.</li>
		<li>Develop a shallow subcapsular pocket that is slightly larger than the valve leaflet while lifting the edge of the incision with one set of forceps and advancing a blunt probe under the renal capsule.</li>
	</ol>
	</li>
	<li>Transplant the aortic valve into the subcapsular pocket.
	<ol>
		<li>Retrieve the aortic leaflet from cold storage and place it in the surgical field.</li>
		<li>While lifting the edge fibrous capsule, advance the aortic leaflet into the subcapsular pocket with blunt forceps. 
		NOTE: Ensure the tissue is far enough away from the incision so that it is firmly secured under the capsule. Care should be taken to avoid damage to the underlying parenchyma or further ripping of the fibrous capsule.</li>
		<li>The incision in the renal capsule can be left open.</li>
	</ol>
	</li>
	<li>Push the kidney gently back to its anatomical position using counter traction applied to the incision edges.</li>
	<li>Close the abdominal incision with a running sterile surgical suture. Close the skin with staples.</li>
	<li>Post-operative care
	<ol>
		<li>Following the operation, place the rat in a clean cage on a heating pad with access to food and water.</li>
		<li>Monitor the animal daily to assess for routine wound healing and signs of pain or distress. Remove the staples after 7-10 days.</li>
	</ol>
	</li>
</ol>
6. Collection of tissue for analysis
<ol>
	<li>At selected endpoints after transplantation, euthanize the animal by exsanguination. Specifically, perform a median laparotomy and transect the abdominal aorta under 5% isoflurane in oxygen.</li>
	<li>Mobilize the kidney and excise it by cutting the renal artery, vein, and ureter with scissors. 
	NOTE: Take care not to grasp the area containing the transplanted leaflet.</li>
	<li>Place the kidney in formalin overnight, embed it in paraffin, and section it for the desired staining. Orient the specimen with the kidney capsule facing anteriorly and the kidney parenchyma facing posteriorly.</li>
</ol>

<ol>
	<li>Van Der Linde, D., 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=Birth+prevalence+of+congenital+heart+disease+worldwide:+A+systematic+review+and+meta-analysis.">Birth prevalence of congenital heart disease worldwide: A systematic review and meta-analysis.</a> Journal of the American College of Cardiology. 58 (21), 2241-2247 (2011).</li><li>Jacobs, J. P., 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=Reoperations+for+pediatric+and+congenital+heart+disease:+An+analysis+of+the+Society+of+Thoracic+Surgeons+(STS)+congenital+heart+surgery+database.">Reoperations for pediatric and congenital heart disease: An analysis of the Society of Thoracic Surgeons (STS) congenital heart surgery database.</a> Seminars in Thoracic and Cardiovascular Surgery: Pediatric Cardiac Surgery Annual. 17 (1), 2-8 (2014).</li><li>Syedain, Z. 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M., Drews, J. D., Breuer, C. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue-engineered+heart+valves:+A+call+for+mechanistic+studies.">Tissue-engineered heart valves: A call for mechanistic studies.</a> Tissue Engineering Part B: Reviews. 24 (3), 240-253 (2018).</li><li>Bernstein, D., 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=Cardiac+growth+after+pediatric+heart+transplantation.">Cardiac growth after pediatric heart transplantation.</a> Circulation. 85 (4), 1433-1439 (1992).</li><li>Delmo Walter, E. 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=Adaptive+growth+and+remodeling+of+transplanted+hearts+in+children.">Adaptive growth and remodeling of transplanted hearts in children.</a> Europeon Journal of Cardiothoracic Surgery. 40 (6), 1374-1383 (2011).</li><li>Simon, P., 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=Growth+of+the+pulmonary+autograft+after+the+Ross+operation+in+childhood.">Growth of the pulmonary autograft after the Ross operation in childhood.</a> Europeon Journal of Cardiothoracic Surgery. 19 (2), 118-121 (2001).</li><li>Oei, F. B. 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F., 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=Prevention+of+allograft+heart+valve+failure+in+a+rat+model.">Prevention of allograft heart valve failure in a rat model.</a> Journal of Thoracic and Cardiovascualr Surgery. 122 (2), 310-317 (2001).</li><li>Legare, J. F., Lee, T. D. G., Creaser, K., Ross, D. 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The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Importance and potential applications 
While mechanical and bioprosthetic heart valves are routinely used in adult patients requiring valve replacement, these valves lack the potential to grow and, therefore, are suboptimal for pediatric patients. Heart valve transplantation is an experimental operation designed to deliver growing heart valve replacements for neonates and infants with congenital heart disease. However, unlike the transplant immunobiology of conventional heart transplants, the transp...

Congenital heart defects are the most common congenital disability in humans, affecting 7 in 1,000 live-born children each year1. Unlike adult patients in which various mechanical and bioprosthetic valves are routinely implanted, pediatric patients currently have no good options for valve replacement. These conventional implants do not have the potential to grow in recipient children. As a result, morbid re-operations are required to exchange the heart valve implants for successively larger versions as the children grow, with affected kids often requiring up to five or more open-heart surgeries in their lifetime2,3. Studies have shown that freedom from intervention or death is significantly poor for infants than older children, with 60% of infants with prosthetic heart valves facing re-operation or death within 3 years of their initial operation4. Therefore, there is an urgent need to deliver a heart valve that can grow and maintain function in pediatric patients.
For decades, attempts to deliver growing heart valve replacements have been centered on tissue engineering and stem cells. However, attempts to translate these valves to the clinic have been unsuccessful thus far5,6,7,8. For addressing this, a heart valve transplantation is proposed as a more creative operation for delivering growing heart valve replacements having the ability to self-repair and avoid thrombogenesis. Instead of transplanting the whole heart, only the heart valve is transplanted and will then grow with the recipient child, similar to conventional heart transplants or a Ross pulmonary autograph9,10,11. Post-operatively, recipient children will receive immunosuppression until the transplanted valve can be exchanged for an adult-sized mechanical prosthetic when the growth of the valve is no longer required. However, the transplant biology of heart valve transplant grafts remains unexplored. Therefore, animal models are needed to study this new type of transplant.
Several rat models have been previously described for heterotopic transplantation of the aortic valve into the abdominal aorta12,13,14,15,16,17,18. However, these models are prohibitively tricky, often requiring trained surgeons to operate successfully. Additionally, they are costly and time-consuming19. A novel rat model was developed to create a simpler animal model for studying the immunobiology of heart valve transplants. Single aortic valve leaflets are excised and inserted into the renal subcapsular space. The kidney is especially suited to study transplant rejection as it is highly vascularized with access to circulating immune cells20,21. While several others have utilized a renal subcapsular model to study the transplant biology of other allograft transplants such as pancreas, liver, kidney, and cornea22,23,24,25,26,27, this is the first description of transplantation of cardiac tissue in this position. Here, the transplantation technique is described, providing a significant step forward in studying the transplant immunology of heart valve transplantation.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.9% Sodium Chlordie, USP</td>
			<td>Baxter</td>
			<td>NDC 0338-0048-04</td>
			<td></td>
		</tr>
		<tr>
			<td>4-0 Polyglactin 910</td>
			<td>Ethicon</td>
			<td>J415H</td>
			<td></td>
		</tr>
		<tr>
			<td>7.5% Povidone-Iodine</td>
			<td>CareFusion</td>
			<td>29904-004</td>
			<td></td>
		</tr>
		<tr>
			<td>70% ETOH</td>
			<td>Fisher Scientific</td>
			<td>BP82031GAL</td>
			<td></td>
		</tr>
		<tr>
			<td>Anesthesia induction chamber</td>
			<td>Harvard Apparatus</td>
			<td>75-2030</td>
			<td>Air-tight inducton chamber for rats</td>
		</tr>
		<tr>
			<td>Anesthesia machine</td>
			<td>Harvard Apparatus</td>
			<td>75-0238</td>
			<td>Mobile Anesthesia System with Passive Scavenging</td>
		</tr>
		<tr>
			<td>Anesthesia Mask</td>
			<td>Harvard Apparatus</td>
			<td>59-8255</td>
			<td>Rat anesthesia mask</td>
		</tr>
		<tr>
			<td>Brown Norway Rats (BN/Crl)</td>
			<td>Charles River</td>
			<td>Strain Code 091</td>
			<td>Male, 5-7 weeks, 100-200 g</td>
		</tr>
		<tr>
			<td>Buprenorphine Hydrochloride, 0.3 mg/mL</td>
			<td>PAR Pharmaceutical</td>
			<td>NDC 42023-179-05</td>
			<td>0.03 mg/kg, administered subcutaneously</td>
		</tr>
		<tr>
			<td>Electric hair clippers</td>
			<td>WAHL</td>
			<td>79434</td>
			<td></td>
		</tr>
		<tr>
			<td>Electric Heating Pad</td>
			<td>Harvard Apparatus</td>
			<td>72-0492</td>
			<td>Maintained at 36-38 &#176;C</td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>Sagent Pharmaceuticals</td>
			<td>NDC 25021-400-10</td>
			<td>100U/100g injection into the left atrium</td>
		</tr>
		<tr>
			<td>Insulin Syringe, 1 mL</td>
			<td>Fisher Scientific</td>
			<td>14-841-33</td>
			<td></td>
		</tr>
		<tr>
			<td>Iris forceps curved</td>
			<td>World Precision Instruments</td>
			<td>15917</td>
			<td></td>
		</tr>
		<tr>
			<td>Iris forceps straight</td>
			<td>World Precision Instruments</td>
			<td>15916</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane, USP</td>
			<td>Piramal Critical Care</td>
			<td>NDC 66794-017-25</td>
			<td>Induced at 5% isoflurance in oxygen and maintained with 3.5% isoflurane in oxygen</td>
		</tr>
		<tr>
			<td>Lewis Rats (LEW/ Crl)</td>
			<td>Charles River</td>
			<td>Strain Code 004</td>
			<td>Male, 5-7 weeks, 100-200 g</td>
		</tr>
		<tr>
			<td>Micro forceps</td>
			<td>World Precision Instruments</td>
			<td>500233</td>
			<td>Dumont #5</td>
		</tr>
		<tr>
			<td>Micro scissors</td>
			<td>World Precision Instruments</td>
			<td>501930</td>
			<td>Spring-loaded Vannas Scissors</td>
		</tr>
		<tr>
			<td>Needle Driver</td>
			<td>World Precision Instruments</td>
			<td>500226</td>
			<td>Ryder Needle Driver</td>
		</tr>
		<tr>
			<td>Operating microscope</td>
			<td>AmScope</td>
			<td>SM-3BZ-80S</td>
			<td>3.5x - 90x Stereo Microscope</td>
		</tr>
		<tr>
			<td>Petri Dish</td>
			<td>Fisher Scientific</td>
			<td>FB0875714</td>
			<td></td>
		</tr>
		<tr>
			<td>Petrolatum ophthalmic ointment</td>
			<td>Dechra</td>
			<td>NDC 17033-211-38</td>
			<td></td>
		</tr>
		<tr>
			<td>Skin staples</td>
			<td>Ethicon</td>
			<td>PXR35</td>
			<td>Proximate 35</td>
		</tr>
		<tr>
			<td>Sterile cotton swabs</td>
			<td>Puritan</td>
			<td>25-806 1WC</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile gauze sponges</td>
			<td>Fisher Scientific</td>
			<td>22-037-902</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Scissors</td>
			<td>World Precision Instruments</td>
			<td>1962C</td>
			<td>Metzenbaum Scissors</td>
		</tr>
		<tr>
			<td>University of Wisconsin Buffer (Servator B)</td>
			<td>S.A.L.F S.p.A.</td>
			<td>6484A1</td>
			<td>Stored at 4 &#176;C</td>
		</tr>
	</tbody>
</table>

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.

A graphical depiction of the experimental design is provided for the rat model (Figure 1). Additionally, an aortic root dissected from the donor&#39;s heart and an individual aortic valve leaflet prepared for implantation is also shown in Figure 2. Next, a representative image of the position of the aortic valve leaflet under the renal capsule for implantation is shown in Figure 3A and after 3, 7, and 28 days within the recipient ra...

Watch this Scientific Journal Video about A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents at JoVE.com

A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents

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-simplified-model-for-heterotopic-heart-valve-transplantation

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JoVE Journal

Medicine

2.2K Views.  Medical University of South Carolina. 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.

Video: A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents

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 ...

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 ...

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 ...

A Modified Cuff Technique for Mouse Cervical Heterotopic Heart Transplantation Model

Cardiac allograft rejection limits the long-term survival of patients after heart transplantation. A mouse heart transplantation model is ideal for investigating the mechanism of cardiac allograft rejection in preclinical studies because of their high homology with human genes. This understanding would help develop unique approaches to improving patients&#39; long-term survival treated with cardiac allografts. In a mouse model, abdominal donor heart implantation is commonly performed with an end-to-end&#160;anastomosis to the recipient&#39;s aorta and inferior vena cava using stitches. In this model, the donor&#39;s heart is implanted by end-to-end anastomosis to the recipient&#39;s carotid artery and jugular vein by the modified-Cuff technique. The transplantation surgery is performed without stitching and thus may increase the survival of the recipient since there is no interference with the blood supply and venous reflux of the lower body. This mouse model would help investigate the mechanisms underlying the immunological and pathological (acute/chronic) rejection of cardiac allografts.

In the present protocol, a mouse heart transplantation model is used for investigating the mechanism of cardiac allograft rejection. In this heterotopic heart transplantation model, operation efficiency is improved, and the survival of cardiac grafts is ensured by a cervical end-to-end anastomosis of heart implantation using a modified Cuff technique.

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

Transcatheter Pulmonary Valve Replacement from Autologous Pericardium with a Self-Expandable Nitinol Stent in an Adult Sheep Model

Transcatheter pulmonary valve replacement has been established as a viable alternative approach for patients suffering from right ventricular outflow tract or bioprosthetic valve dysfunction, with excellent early and late clinical outcomes. However, clinical challenges such as stented heart valve deterioration, coronary occlusion, endocarditis, and other complications must be addressed for lifetime application, particularly in pediatric patients. To facilitate the development of a lifelong solution for patients, transcatheter autologous pulmonary valve replacement was performed in an adult sheep model. The autologous pericardium was harvested from the sheep via left anterolateral minithoracotomy under general anesthesia with ventilation. The pericardium was placed on a 3D shaping heart valve model for non-toxic cross-linking for 2 days and 21 h. Intracardiac echocardiography (ICE) and angiography were performed to assess the position, morphology, function, and dimensions of the native pulmonary valve (NPV). After trimming, the crosslinked pericardium was sewn onto a self-expandable Nitinol stent and crimped into a self-designed delivery system. The autologous pulmonary valve (APV) was implanted at the NPV position via left jugular vein catheterization. ICE and angiography were repeated to evaluate the position, morphology, function, and dimensions of the APV. An APV was successfully implanted in&#160;sheep J. In this paper, sheep J was selected to obtain representative results. A 30 mm APV with a Nitinol stent was accurately implanted at the NPV position without any significant hemodynamic change. There was no paravalvular leak, no new pulmonary valve insufficiency, or stented pulmonary valve migration. This study demonstrated the feasibility and safety, in a long-time follow-up, of developing an APV for implantation at the NPV position with a self-expandable Nitinol stent via jugular vein catheterization in an adult sheep model.

This study demonstrates the feasibility and safety of developing an autologous pulmonary valve for implantation at the native pulmonary valve position by using a self-expandable Nitinol stent in an adult sheep model. This is a step toward developing transcatheter pulmonary valve replacement for patients with right ventricular outflow tract dysfunction.

Transcatheter pulmonary valve replacement has been established as a viable alternative approach for patients suffering from right ventricular outflow ...

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.

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 ...