Name: a simplified model for heterotopic heart valve transplantation in rodents
Uploaded: 2021-09-21T14:30:00.000+00:00
Duration: 6 min 33 s
Description: Watch this Scientific Journal Video about A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents at JoVE.com

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. 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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><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

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

Research

JoVE Journal

Medicine

2.3K 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

Heterotopic Heart Transplantation in Mice

The mouse heterotopic heart transplantation has been used widely since it was introduced by Drs. Corry and Russell in 1973. It is particularly valuable for studying rejection and immune response now that newer transgenic and gene knockout mice are available, and a large number of immunologic reagents have been developed. The heart transplant model is less stringent than the skin transplant models, although technically more challenging. We have developed a modified technique and have completed over 1000 successful cases of heterotopic heart transplantation in mice. When making anastomosis of the ascending aorta and abdominal aorta, two stay sutures are placed at the proximal and distal apexes of recipient abdominal aorta with the donor s ascending aorta, then using 11-0 suture for anastomosis on both side of aorta with continuing sutures. The stay sutures make the anastomosis easier and 11-0 is an ideal suture size to avoid bleeding and thrombosis.
When making anastomosis of pulmonary artery and inferior vena cava, two stay sutures are made at the proximal apex and distal apex of the recipient s inferior vena cava with the donor s pulmonary artery. The left wall of the inferior vena cava and donor s pulmonary artery is closed with continuing sutures in the inside of the inferior vena cava after, one knot with the proximal apex stay suture the right wall of the inferior vena cava and the donor s pulmonary artery are closed with continuing sutures outside the inferior vena cave with 10-0 sutures. This method is easier to perform because anastomosis is made just on the one side of the inferior vena cava and 10-0 sutures is the right size to avoid bleeding and thrombosis. In this article, we provide details of the technique to supplement the video.

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.

The mouse heterotopic heart transplantation has been used widely since it was introduced by Drs. Corry and Russell in 1973. It is particularly valuable ...

Biology

Murine Cervical Heart Transplantation Model Using a Modified Cuff Technique

Mouse models are of special interest in research since a wide variety of monoclonal antibodies and commercially defined inbred and knockout strains are available to perform mechanistic in vivo studies. While heart transplantation models using a suture technique were first successfully developed in rats, the translation into an equally widespread used murine equivalent was never achieved due the technical complexity of the microsurgical procedure. In contrast, non-suture cuff techniques, also developed initially in rats, were successfully adapted for use in mice1-3. This technique for revascularization involves two major steps I) everting the recipient vessel over a polyethylene cuff; II) pulling the donor vessel over the formerly everted recipient vessel and holding it in place with a circumferential tie. This ensures a continuity of the endothelial layer, short operating time and very high patency rates4.
Using this technique for vascular anastomosis we performed more than 1,000 cervical heart transplants with an overall success rate of 95%. For arterial inflow the common carotid artery and the proximal aortic arch were anastomosed resulting in a retrograde perfusion of the transplanted heart. For venous drainage the pulmonary artery of the graft was anastomosed with the external jugular vein of the recipient5.
Herein, we provide additional details of this technique to supplement the video.

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.

Mouse models are of special interest in research since a wide variety of monoclonal antibodies and commercially defined inbred and knockout strains are ...

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 ...

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

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

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

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

Heterotopic Cervical Heart Transplantation in Mice

The heterotopic cervical heart transplantation in mice is a valuable tool in transplant and cardiovascular research. The cuff technique greatly simplifies this model by avoiding challenging suture anastomoses of small vessels thereby reducing warm ischemia time. In comparison to abdominal graft implantation the cervical model is less invasive and the implanted graft is easily accessible for further follow-up examinations. Anastomoses are performed by pulling the ascending aorta of the graft over the cuff with the recipient&#8217;s common carotid artery and by pulling the main pulmonary artery over the cuff with the external jugular vein. Selection of appropriate cuff size and complete mobilization of the vessels are important for successful revascularization. Ischemia-reperfusion (I/R) injury can be minimized by perfusing the graft with a cardioplegic solution and by hypothermia. In this article, we provide technical details for a simplified and improved cuff technique, which should allow surgeons with basic microsurgical skills to perform the procedure with a high success rate.

The cuff technique shortens and facilitates the model of heterotopic cervical heart transplantation in mice by avoiding technically challenging suture anastomoses of small vessels. The technical details shown in this video paper should allow researches to establish this model in their laboratories.

The heterotopic cervical heart transplantation in mice is a valuable tool in transplant and cardiovascular research. The cuff technique greatly simplifies ...

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

Heterotopic heart transplantation in rats has been a commonly used model for diverse immunological studies for more than 50 years. Several modifications have been reported since the first description in 1964. After 30 years of performing heterotopic heart transplantation in rats, we have developed a simplified surgical approach, which can be easily taught and performed without further surgical training or background.
After dissection of the ascending aorta and the pulmonary artery and ligation of superior and inferior caval and pulmonary veins, the donor heart is harvested and subsequently perfused with ice-cold saline solution supplemented with heparin. After clamping and incising the recipient abdominal vessels, the donor ascending aorta and pulmonary artery are anastomosed to the recipient abdominal aorta and inferior vena cava, respectively, using continuous running sutures.
Depending on different donor-recipient combinations, this model allows analyses of either acute or chronic rejection of allografts. The immunological significance of this model is further enhanced by a novel approach of in-ear injection of vital cardiac muscle cells and subsequent analysis of draining cervical lymphatic tissue.

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.

Heterotopic heart transplantation in rats has been a commonly used model for diverse immunological studies for more than 50 years. Several modifications ...

Immunology and Infection

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 ...

Blood Circuit Reconstruction in an Abdominal Mouse Heart Transplantation Model

The surgical technique of heterotopic abdominal heart transplantation in mice is a standard model for research in transplantation immunology. Here, the established technique for a modified blood circuit reconstruction in a heterotopic abdominal heart transplantation model is presented. This method uses the intrathoracic inferior vena cava (IIVC) instead of the pulmonary artery of the donor heart for the anastomosis to the inferior vena cava of the recipient. It is facilitating and improving success rates for abdominal heart transplantation in mice.

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

The surgical technique of heterotopic abdominal heart transplantation in mice is a standard model for research in transplantation immunology. Here, the ...

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 ...

A Heterotopic Rat Heart Transplantation Model using Circulatory Death Donor Hearts

The objective of this protocol is to set up a rat heterotopic heart transplantation model with donation after circulatory death (DCD) donor hearts. There are two setups for this protocol: heart donor setup and recipient setup. In the heart donor setup, Sprague Dawley rats are anesthetized, endotracheally intubated, and ventilated. The right carotid artery is cannulated to deliver heparin and the paralytic agent vecuronium-bromide. The DCD process is initiated by terminating the ventilation. After 20 min, the heart is exposed and the aorta distal to the brachiocephalic branch is clamped. At 25 min from terminating the ventilator, ice-cold University of Wisconsin (UW) solution is perfused through the carotid catheter to flush the heart. The heart is procured by dividing the aorta, pulmonary artery, venae cavae, and pulmonary veins and stored in UW solution for implantation. In the recipient setup, the Lewis rat is anesthetized with isoflurane. Slow-release buprenorphine is administered subcutaneously to facilitate a smooth postoperative recovery. Through a midline abdominal incision, the infra-renal aorta and the inferior vena cava are isolated and clamped with an atraumatic vascular clamp. The donor heart aorta and pulmonary artery are sutured to the recipient abdominal aorta and vena cava, respectively, with a running 8-0 Prolene. The vascular clamp is removed to reperfuse the heart. The abdominal wall is closed and the rat is recovered. After a set interval (24 h to 2 weeks), the recipient rat is anesthetized, the transplanted heart is exposed, and a balloon-tip-catheter is inserted into the left ventricle via the apex to record developed pressure and dP/dt using a data acquisition system. The heart tissue is collected for histology, immunology, or molecular analysis. A successful DCD donor rat heart transplantation model will allow further studies on the cardioprotective approaches to improve heart transplantation outcomes from DCD donors.

Here, we describe the preparation and technical details of murine heterotopic heart transplantation utilizing a circulatory death donor heart.

The objective of this protocol is to set up a rat heterotopic heart transplantation model with donation after circulatory death (DCD) donor hearts. There ...