This work was supported by the National Health and Medical Research Council (NHMRC) of Australia Project Grant 1029205, by the Microsearch Foundation of Australia, and by the Myee Codrington Medical Research Foundation.

Prior to commencement of experiments, obtain approval from the relevant institution&#8217;s Animal Care Ethics Committee for the planned experiments. Maintain mice in the accordance with the requirements of your institution. The following protocol has been approved by Sydney University and Royal Prince Alfred Hospital committees.
Prior to commencing surgery, all instruments should be cleaned thoroughly and sterilized by soaking in 80% ethanol. Some institutions stipulate autoclaving however this can damage fine microsurgical instruments over the long term. &#160;Where possible use sterile disposable equipment.
1. Anesthesia
<ol>
	<li>Anesthetize mouse with isoflurane in a sealed container then place it supine on an operating board, quickly connecting anesthetic nose cone. Test adequacy of anesthesia by pinching the hind foot to ensure there is no reflex.</li>
	<li>Shave the skin with a surgical blade and sterilize with 80% ethanol. In addition, 2% chlorhexidine in 70% isopropyl alcohol can be used for skin sterilization.&#160;During induction of anesthesia the isoflurane concentration is 3% but reduce this to 1-1.5% for maintenance. Make minor adjustments to this concentration to maintain regular breathing and heart rate. Maintain temperature of the animal on a heating pad.</li>
</ol>
2. Donor Operation
<ol>
	<li>Make a thoracotomy to expose the heart and vessels by cutting the chest through both sides of the rib cage from the rib edge up to the axilla followed by a transverse cut at the level of the xiphisternum to form a chest wall flap. Lift this flap up beside the head and pin it to the operating board. Tear off pericardium to expose the heart and vessels.</li>
	<li>Lift the inferior vena cava with forceps in one hand and with the other inject 1 ml cold heparinized saline proximal to heart through the IVC, then put a small artery clamp on the IVC to prevent flow of perfusion solution back through the needle hole.</li>
	<li>Using gauze and a cotton bud, retract the heart downward to expose the ascending aorta and pulmonary artery. Pass one blade of a pair of microscissors through the channel (transverse sinus) posterior to the bundle of aorta and pulmonary artery and cut the aorta and pulmonary artery together as far distally as possible to ensure sufficient length for anastomosis.</li>
	<li>Tie and divide the IVC, right superior vena cava (SVC), left SVC and pulmonary veins using 6-0 silk thread. Tie the IVC and right SVC separately then place a single thread posterior to the heart to encircle the left SVC together with pulmonary veins and tie it. Harvest the heart from the donor site by cutting the vessels distal to the ties, then store it in cold sterile saline at 4 &#176;C until transplantation. This results in death of the donor by exsanguination.</li>
</ol>
3. Recipient Operation
<ol>
	<li>Anesthetize the recipient as above (section 1). Carefully shave abdomen to avoid irritation,&#160;then make a laparotomy by midline incision from pubis to xiphisternum and retract using paper clips bent to form retractors. Wrap the bowel in warm sterile saline soaked gauze and retract to the upper right of the abdomen.</li>
	<li>To expose the infra-renal aorta and IVC, free the bundled segments of aorta and IVC from the left renal artery and vein to the iliac bifurcation by dividing them from the lumbar vessels using a cautery device. Take care that the cautery is at the correct temperature and used for sufficient time to divide and seal the vessel ends.</li>
	<li>Apply small atraumatic vessel clamps to the aorta and IVC segments distally and proximally. Make an incision in the front wall of the aorta by first puncturing a hole with a 30 G&#160;needle; then cut an incision vertically with microscissors to match the size of the donor aorta. Flush the aorta lumen with heparinized saline to remove any blood clots.</li>
	<li>Bring the donor heart to the recipient site covered with gauze soaked in cold saline and place it to the right side of the abdomen. Make sure the donor aorta is positioned next to the incision on the recipient aorta and the donor pulmonary artery positioned next to the recipient IVC.</li>
	<li>Anastomose the donor aorta end-to-side to the recipient aorta using running 10-0 nylon sutures starting from the proximal corner and suturing along the left side first until reaching the distal corner, at which stage the animal is rotated through 180&#176;. Gently move the donor heart to the left side of the abdomen to expose the right side of the aorta and continue suturing through the right side of the aortic wall from the distal end to the proximal end. Before closing the aortic anastomosis, gently flush the lumen with heparinized saline to remove any clot and air.</li>
	<li>Anastomose the donor pulmonary artery (PA) end-to-side with the recipient IVC. Make an incision vertically with microscissors in the front wall of the IVC at a site in accordance with the artery anastomosis. Anastomose the donor PA to recipient IVC with running 10-0 nylon sutures starting from the distal end of the left wall within the lumen of the IVC. After reaching the proximal end, continue sutures along the front right side wall up to the distal end to complete the anastomosis. Before closing the anastomosis, gently flush the lumen to remove any clots and air.</li>
	<li>Before releasing the vessel clamps, place pieces of Gelfoam around the anastomosis sites, and apply gentle pressure with a cotton applicator until hemostasis is achieved. At the time of revascularization, release the distal clamp first, followed by the proximal clamp.</li>
	<li>After revascularization, apply warm saline at 37 &#176;C to the graft externally to assist its recovery. The graft usually starts fibrillation immediately and reverts spontaneously to sinus rhythm within a few minutes. Inject 0.6 ml warm saline subcutaneously to maintain hydration of the recipient. Inject buprenorphine subcutaneously for analgesia prior to completion of surgery</li>
	<li>Close the abdominal wound with one 5-0 absorbable running suture for&#160;both the layers. Start by completing the inner layer and continue along the skin.</li>
</ol>
4. Recovery and Graft Monitoring
<ol>
	<li>Inject ampicillin for prophylaxis of infection and place the recipient on a heating pad at 37 &#176;C for recovery. Most animals recover rapidly and are usually drinking and often eating within 3 hr. If the mouse shows signs of distress, examine closely to determine cause. If no obvious cause, treat with buprenorphine and monitor closely. Consult a veterinarian if symptoms are severe or persist more than 8 hours and consider euthanasia. If necessary, give continuing 12&#160;hourly injections of buprenorphine until symptoms resolve.&#160;If the mouse shows signs of distress after 8 hr, give continuing 12 hourly injections of buprenorphine until they resolve. Consult the veterinarian if symptoms continue for more than 48 hr and euthanize if necessary.</li>
	<li>Monitor graft heart beat by direct abdominal palpation and record the strength of beat as ++++ for a healthy graft to + for weak beat due to advanced rejection and &#8211; as non beating due to complete rejection of the graft. Monitor mice daily for the first 10 days, then 3x per week for the duration of the experiment.</li>
</ol>

<ol>
	<li>Aramaki, O., 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=Interleukin-10+but+not+transforming+growth+factor-beta+is+essential+for+generation+and+suppressor+function+of+regulatory+cells+induced+by+intratracheal+delivery+of+alloantigen.">Interleukin-10 but not transforming growth factor-beta is essential for generation and suppressor function of regulatory cells induced by intratracheal delivery of alloantigen.</a> Transplantation. 79, 568-576 (2005).</li><li>Chen, R. H., Bushell, A., Fuggle, S. V., Wood, K. J., Morris, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+of+granzyme+A+and+perforin+in+mouse+heart+transplants+immunosuppressed+with+donor-specific+transfusion+and+anti-CD4+monoclonal+antibody.">Expression of granzyme A and perforin in mouse heart transplants immunosuppressed with donor-specific transfusion and anti-CD4 monoclonal antibody.</a> Transplantation. 61, 625-629 (1996).</li><li>Poulin, L. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interleukin-9+promotes+eosinophilic+rejection+of+mouse+heart+allografts.">Interleukin-9 promotes eosinophilic rejection of mouse heart allografts.</a> Transplantation. 76, 572-577 (2003).</li><li>Ono, K., Lindsey, E. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improved+technique+of+heart+transplantation+in+rats.">Improved technique of heart transplantation in rats.</a> J. Thorac. Cardiovasc. Surg. 57, 225-229 (1969).</li><li>Corry, R. J., Winn, H. J., Russell, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Primarily+vascularized+allografts+of+heart+in+mice.">Primarily vascularized allografts of heart in mice.</a> Transplantation. 16, 343-350 (1973).</li><li>Larsen, C. P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CD40-gp39+interactions+play+a+critical+role+during+allograft+rejection.+Suppression+of+allograft+rejection+by+blockade+of+the+CD40-gp39+pathway.">CD40-gp39 interactions play a critical role during allograft rejection. Suppression of allograft rejection by blockade of the CD40-gp39 pathway.</a> Transplantation. 61, 4-9 (1996).</li><li>Saitovitch, D., Bushell, A., Mabbs, D. W., Morris, P. J., Wood, K. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinetics+of+induction+of+transplantation+tolerance+with+a+nondepleting+anti-Cd4+monoclonal+antibody+and+donor-specific+transfusion+before+transplantation.+A+critical+period+of+time+is+required+for+development+of+immunological+unresponsiveness.">Kinetics of induction of transplantation tolerance with a nondepleting anti-Cd4 monoclonal antibody and donor-specific transfusion before transplantation. A critical period of time is required for development of immunological unresponsiveness.</a> Transplantation. 61, 1642-1647 (1996).</li><li>Callaghan, C. J., 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=Regulation+of+allograft+survival+by+inhibitory+FcgammaRIIb+signaling.">Regulation of allograft survival by inhibitory FcgammaRIIb signaling.</a> J. Immunol. 189, 5694-5702 (2012).</li><li>Qian, S., 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=Impact+of+donor+MHC+Class+I+or+Class+II+antigen+deficiency+on+first-+and+second-set+rejection+of+mouse+heart+or+liver+allografts.">Impact of donor MHC Class I or Class II antigen deficiency on first- and second-set rejection of mouse heart or liver allografts.</a> Immunology. 88, 124-129 (1996).</li><li>Wang, C., 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=Spontaneous+acceptance+of+mouse+kidney+allografts+is+associated+with+increased+Foxp3+expression+and+differences+in+the+B+and+T+cell+compartments.">Spontaneous acceptance of mouse kidney allografts is associated with increased Foxp3 expression and differences in the B and T cell compartments.</a> Transpl. Immunol. 24, 149-156 (2011).</li><li>Liu, F., Kang, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heterotopic+Heart+Transplantation+in+Mice.">Heterotopic Heart Transplantation in Mice.</a> J. Vis. Exp. , (2007).</li><li>Mottram, P. L., 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=Electrocardiographic+monitoring+of+cardiac+transplants+in+mice.">Electrocardiographic monitoring of cardiac transplants in mice.</a> Cardiovasc. Res. 22, 315-321 (1988).</li></ol>

The authors have nothing to disclose.

Mouse heart transplantation is a challenging microsurgical method that requires considerable surgical skill to master. The most challenging aspect is the small diameter of the vessels. In addition, it is necessary to limit the recipient operation time and bleeding. The technique for mouse heart transplantation was first described by Corry et al. in 1973 and subsequently by Mottram et al12. Our modifications include the following points. Firstly, immediate perfusion of the donor heart with col...

Animal models of organ transplantation can provide valuable information for improving treatment of clinical transplant patients. Mouse models are particularly useful for studying the immune mechanism of organ transplant rejection or acceptance due to the wide range of genetically modified mice and reagents specific for mice that are not available for other animal models1-3. A challenge with mouse models of transplantation is the small size of the donors and recipients which requires considerable technical skill to achieve satisfactory outcomes.
A technique was first described for heterotopic transplantation of hearts in rats4 which was subsequently adapted for mouse heart transplantation by Corry et al5. This technique involved preparing the recipient prior to the donor operation and no perfusion of the donor heart, both of which are likely to compromise survival of the transplanted heart or the recipient. The procedure has been widely used as originally described to examine the mechanisms of transplant rejection and tolerance6-8. Others have adapted the original rat heart transplant procedure of Ono and Lindsey for heart transplants in mice9,10. More recently, a technique was published for mouse heart transplantation that overcame some of the problems associated with the method of Corry et al11. The protocol described here incorporates our modifications, based on the method of Mottram et al12, which include: in situ perfusion with cold heparinized saline immediately after thoracotomy and performing the donor operation prior to the recipient operation to minimize recipient operation time. In addition, we use small atraumatic vessel clamps instead of 6-0 silk ties. Although vessel clamps have the disadvantage of taking more space they are easier to control than ties, which should not be too tight or loose and are less straightforward to remove than clamps. Our method uses a single running suture for vessel anastomosis, although in the initial learning stages an alternative is to use stay sutures at the proximal and distal corners to ensure evenness of sutures and thus patency of the anastomosis.

<table><tbody><tr><td>Operating microscope&amp;nbsp;</td><td>Leica, Heerbrugg, Switzerland</td><td>M651</td><td>10-25X magnification</td></tr><tr><td>Anesthetic machine&amp;nbsp;</td><td>Vet Quip Pty Ltd, Sydney, Australia</td><td>Vett3</td><td>Capable of delivering a mixture of isoflurane and oxygen in air&amp;nbsp;</td></tr><tr><td>Operating board&amp;nbsp;</td><td>Hardware store or office supplier</td><td></td><td>Dense cork or synthetic capable of taking pins</td></tr><tr><td>Heparinized saline (5 U/ml, 4 &amp;deg;C)</td><td>Pfizer, USA</td><td>FW25</td><td>In 1 ml syringe with 23 G needle on ice</td></tr><tr><td>Normal saline (37 &amp;deg;C)</td><td>AstraZeneca Pty Ltd, Australia</td><td>4538</td><td>In 1 ml syringe with 23 G needle on warm pad</td></tr><tr><td>Sutures 10- nylon, 5-0 Vicryl</td><td>Ethicon, Inc. NJ, USA</td><td>2870G/J421H</td><td></td></tr><tr><td>Buprenorphine (0.05 mg/kg in 0.1 ml saline)</td><td>Reckitt Benckiser, Sydney, Australia</td><td></td><td>In 1 ml syringe with 23 G needle on ice</td></tr><tr><td>Ampcillin (5 mg/kg in 0.1 ml saline)</td><td>Aspen, Sydney, Australia</td><td></td><td>In 1 ml syringe with 23 G needle on ice</td></tr><tr><td>Gel Foam</td><td>Pharmacia &amp;amp; Upjohn Co. USA</td><td>801289304</td><td>Cut into small pieces</td></tr><tr><td>High temperature cautery device</td><td>Medtronic, USA</td><td>8444000</td><td></td></tr><tr><td>Heating Pad/Right Temp</td><td>Kent Scientific Corporation, Turrington, CT 06790</td><td></td><td></td></tr><tr><td>&lt;strong&gt;Microsurgery instruments:&lt;/strong&gt;</td><td>Shanghai Medical Instruments Co. Ltd.,&amp;nbsp;&amp;nbsp;</td><td></td><td></td></tr><tr><td>Microneedleholders</td><td>Shanghai, China</td><td>WT2020</td><td></td></tr><tr><td>Microscissors</td><td>&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; &quot;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; &quot;</td><td>WT1020</td><td></td></tr><tr><td>Microforceps (straight tip)</td><td>&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; &quot;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; &quot;</td><td>WA3010</td><td></td></tr><tr><td>Microforceps (curved tip)</td><td>&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; &quot;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; &quot;</td><td>WA3020</td><td></td></tr><tr><td>Micromosquito clamps (1 pair)</td><td>&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; &quot;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; &quot;</td><td>W40350</td><td></td></tr><tr><td>Microvessel atraumatic clamps (1 pair)</td><td>&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; &quot;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; &quot;</td><td>W40130/W40150</td><td></td></tr></tbody></table>

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.

After an initial training period, 227 cases of mouse heterotopic heart transplantation in our group were analyzed. The success rate within the first 24 hr was 90.3% and at 48 hr it was 86.8%. Of the 30 (13.2%) failures within 48 hr, 5 (2.2%) experienced hind limb paralysis and had to be euthanized, 10 (4.4%) had a non-beating heart due to graft ischemia injury and/or thrombosis, while 15 (6.6%) died within 48 hr. Some experimental graft survivals are shown in Table 1 with a variety of strain combinations...

Watch this Scientific Journal Video about A Modified Method for Heterotopic Mouse Heart Transplantion at JoVE.com

A Modified Method for Heterotopic Mouse Heart Transplantion

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 ...

a-modified-method-for-heterotopic-mouse-heart-transplantion

Universidad de Cartagena UDEC

Research

JoVE Journal

Medicine

12.0K Views.  University of Sydney. 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.

Video: A Modified Method for Heterotopic Mouse Heart Transplantion

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 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 ...

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

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

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

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

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

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 ...

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 ...

A Modified Murine Heterotopic Heart Transplant Protocol Matching Contemporary Standards of Aseptic Technique, Anesthesia, and Analgesia

The development of experimental models of cardiac transplantation in animals has contributed to many advances in the fields of immunology and solid organ transplantation. While the heterotopic vascularized murine cardiac transplantation model was initially utilized in studies of graft rejection using combinations of mismatched inbred mouse strains, access to genetically modified strains and therapeutic modalities can provide powerful new preclinical insights. Fundamentally, the surgical methodology for this technique has not changed since its development, especially with respect to important factors such as aseptic technique, anesthesia, and analgesia, which make material impacts on postsurgical morbidity and mortality. Additionally, improvements in perioperative management are expected to provide improvements in both animal welfare and experimental outcomes. This paper reports upon a protocol developed in collaboration with a subject matter expert in veterinary anesthesia and describes the surgical technique with an emphasis on perioperative management. Additionally, we discuss the implications of these refinements and provide details on troubleshooting critical surgical steps for this procedure.

The present paper describes a modified technique for heterotopic vascularized cardiac transplantation with updated aseptic technique, analgesia, and anesthesia.

The development of experimental models of cardiac transplantation in animals has contributed to many advances in the fields of immunology and solid organ ...

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 ...