Name: a novel microsurgical model for heterotopic, en bloc chest wall, thymus, and heart transplantation in mice
Uploaded: 2016-01-23T14:00:00.000+00:00
Duration: 12 min 24 s
Description: Watch this Scientific Journal Video about A Novel Microsurgical Model for Heterotopic, En Bloc Chest Wall, Thymus, and Heart Transplantation in Mice at JoVE.com

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

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

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

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

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

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


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

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

Exploration of novel strategies in organ transplantation to prolong allograft survival and minimizing the need for long-term maintenance immunosuppression must be pursued. Employing vascularized bone marrow transplantation and co-transplantation of the thymus have shown promise in this regard in various animal models.1-11 Vascularized bone marrow transplantation allows for the uninterrupted transfer of donor bone marrow cells within the preserved donor microenvironment, and the incorporation of thymus tissue with vascularized bone marrow transplantation has shown to increase T-cell chimerism ultimately playing a supportive role in the induction of immune regulation. The combination of solid organ and vascularized composite allotransplantation can uniquely combine these strategies in the form of a novel transplant model. Murine models serve as an excellent paradigm to explore the mechanisms of acute and chronic rejection, chimerism, and tolerance induction, thus providing the foundation to propagate superior allograft survival strategies for larger animal models and future clinical application. Herein, we developed a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique. The experience in syngeneic and allogeneic transplant settings is described for future broader immunological investigations via an instructional manuscript and video supplement.

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

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

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

To study combined solid organ and vascularized composite allotransplantation, we describe a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique.

Exploration of novel strategies in organ transplantation to prolong allograft survival and minimizing the need for long-term maintenance immunosuppression ...

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Research

JoVE Journal

Medicine

9.2K Views.  Johns Hopkins University School of Medicine. To study combined solid organ and vascularized composite allotransplantation, we describe a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique.

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

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 ...

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 ...

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 ...

Optimization of the Cuff Technique for Murine Heart Transplantation

Murine cardiac transplantation has been performed for more than 40 years. With advancements in microsurgery, certain new techniques have been used to improve surgical efficiency. In our lab, we have optimized the cuff technique with two major steps. First, we used the inner tube technique to insert a temporary inner tube into the external jugular vein and carotid artery blood vessel to facilitate eversion of the vessel over the cuff. Second, we performed complete heterotopic cardiac transplantation through the collaboration of two experienced surgeons. These modifications effectively reduced the operation time to 25 minutes, with a success rate of 95%. In this report, we describe these procedures in detail and provide a supplemental video. We believe that this report on the improved cuff technique will offer practical guidance for murine heterotopic heart transplantation and will enhance the utility of this mouse model for basic research.

We introduce an inner tube approach to the cuff technique for mouse cervical heterotopic heart transplantation to help evert the vessel over the cuff. We found that cooperation between two experienced surgeons markedly shortens the operation time.

Murine cardiac transplantation has been performed for more than 40 years. With advancements in microsurgery, certain new techniques have been used to ...

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 ...

Mouse Heterotopic Cervical Cardiac Transplantation Utilizing Vascular Cuffs

Murine models of cardiac transplantation are frequently utilized to study ischemia-reperfusion injury, innate and adaptive immune responses after transplantation, and the impact of immunomodulatory therapies on graft rejection. Heterotopic cervical heart transplantation in mice was first described in 1991 using sutured anastomoses and subsequently modified to include cuff techniques. This modification allowed for improved success rates, and since then, there have been multiple reports that have proposed further technical improvements. However, translation into more widespread utilization remains limited due to the technical difficulty associated with graft anastomoses, which requires precision to achieve adequate length and caliber of the cuffs to avoid vascular anastomotic twisting or excessive tension, which can result in damage to the graft. The present protocol describes a modified technique for performing heterotopic cervical cardiac transplantation in mice which involves cuff placement on the recipient's common carotid artery and the donor's pulmonary artery in alignment with the direction of the blood flow.

Mouse cardiac transplantation models represent valuable research tools for studying transplantation immunology. The present protocol details mouse heterotopic cervical cardiac transplantation that involves the placement of cuffs on the recipient's common carotid artery and the donor's pulmonary artery trunk to allow for laminar blood flow.

Murine models of cardiac transplantation are frequently utilized to study ischemia-reperfusion injury, innate and adaptive immune responses after ...

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 ...