Name: A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents (Video) | JoVE Journal
Uploaded: 2021-09-21T14:30:00.000+00:00
Duration: 6 min 33 s
Description: Learn a simplified heterotopic heart valve transplantation method in rodents using renal subcapsular implantation of aortic valve leaflets. This technique enables researchers to study heart valve immunobiology and alloreactivity effectively.

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5 minutes

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>

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</ol>

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><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 &amp;deg;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 &amp;deg;C</td></tr></tbody></table>

a simplified model for heterotopic heart valve transplantation in rodents

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'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 rat (<...

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

Research

JoVE Journal

Medicine

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

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.

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

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

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.

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

We describe a model of heterotopic abdominal heart transplantation in rats, implying modifications of current strategies, which lead to a simplified surgical approach. Additionally, we describe a novel rejection model by in-ear injection of vital cardiac muscle cells, allowing further transplant immunological analyses in rats.

An Immunological Model for Heterotopic Heart and Cardiac Muscle Cell Transplantation in Rats

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

This study demonstrates a reproducible heterotopic abdominal heart transplantation technique in rats that beginners can learn and perform. Additionally, a novel aortic regurgitation model in rats is generated by performing heterotopic abdominal heart transplantation and damaging the donor's aortic valve using a guidewire after harvesting.

Modified Heterotopic Abdominal Heart Transplantation and a Novel Aortic Regurgitation Model in Rats

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 simplified heterotopic heart valve transplantation method in rodents using renal subcapsular implantation of aortic valve leaflets. This technique enables researchers to study heart valve immunobiology and alloreactivity effectively.

A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents

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