Name: A Modified Method for Heterotopic Mouse Heart Transplantion (Video) | JoVE Journal
Uploaded: 2014-06-23T19:00:00.000+00:00
Duration: 11 min 29 s
Description: Learn a modified microsurgical technique for heterotopic heart transplantation in mice using abdominal vessel anastomosis. This method enables researchers to study transplant rejection mechanisms and strain compatibility in cardiovascular research.

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5 minutes

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/pubmed?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%5BTitle%5D)+AND+%22Transplantation%22%5BJournal%5D)">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/pubmed?term=((Expression+of+granzyme+A+and+perforin+in+mouse+heart+transplants+immunosuppressed+with+donor-specific+transfusion+and+anti-CD4+monoclonal+antibody%5BTitle%5D)+AND+%22Transplantation%22%5BJournal%5D)">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/pubmed?term=((Interleukin-9+promotes+eosinophilic+rejection+of+mouse+heart+allografts%5BTitle%5D)+AND+%22Transplantation%22%5BJournal%5D)">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/pubmed?term=((Improved+technique+of+heart+transplantation+in+rats%5BTitle%5D)+AND+%22J.+Thorac.+Cardiovasc.+Surg%22%5BJournal%5D)">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/pubmed?term=((Primarily+vascularized+allografts+of+heart+in+mice%5BTitle%5D)+AND+%22Transplantation%22%5BJournal%5D)">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/pubmed?term=((CD40-gp39+interactions+play+a+critical+role+during+allograft+rejection.+Suppression+of+allograft+rejection+by+blockade+of+the+CD40-gp39+pathway%5BTitle%5D)+AND+%22Transplantation%22%5BJournal%5D)">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/pubmed?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%5BTitle%5D)+AND+%22Transplantation%22%5BJournal%5D)">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/pubmed?term=((Regulation+of+allograft+survival+by+inhibitory+FcgammaRIIb+signaling%5BTitle%5D)+AND+%22J.+Immunol%22%5BJournal%5D)">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/pubmed?term=((Impact+of+donor+MHC+Class+I+or+Class+II+antigen+deficiency+on+first-+and+second-set+rejection+of+mouse+heart+or+liver+allografts%5BTitle%5D)+AND+%22Immunology%22%5BJournal%5D)">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/pubmed?term=((Spontaneous+acceptance+of+mouse+kidney+allografts+is+associated+with+increased+Foxp3+expression+and+differences+in+the+B+and+T+cell+compartments%5BTitle%5D)+AND+%22Transpl.+Immunol%22%5BJournal%5D)">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/pubmed?term=((Heterotopic+Heart+Transplantation+in+Mice%5BTitle%5D)+AND+%22J.+Vis.+Exp%22%5BJournal%5D)">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/pubmed?term=((Electrocardiographic+monitoring+of+cardiac+transplants+in+mice%5BTitle%5D)+AND+%22Cardiovasc.+Res%22%5BJournal%5D)">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

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...

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 ...

The overall goal of this procedure is to transplant a heart from a donor to a recipient mouse. This is accomplished by first removing the heart from the donor thorax and storing it in cold saline. The next steps are to transplant the heart into the abdomen of the recipient and then monitor the transplanted heartbeat.Finally, transplant rejection is confirmed by visual inspection and pathology analysis. Ultimately, results show that hearts transplanted between unrelated strains of mice are rejected while hearts transplanted between the same strain are accepted. Long term Video demonstration of this measured is critical at the microsurgical step are difficult to learn because of the very small size of donor and the recipient animals.The complicated surgical procedure involved and the need for boost accuracy and the speed in performing the recipient operations Always make certain that all the instruments are thoroughly cleaned and sterilized in an 80%ethanol bath. Use of disposable sterile equipment is highly advisable. After anesthetizing the donor mouse in an isof fluorine chamber position, its supine on the operating board.Then quickly connect the nose cone, delivering anesthetic. Be sure to test the quality of the anesthesia with a hind foot pinch. If there is no flinch response, commence with shaving the skin using a surgical blade.Then sterilize the skin with three alternating wipes of 2%Chlorhexidine in 70%isopropyl alcohol throughout the surgery. Visually monitor the animal's breathing and heartbeat. Adjust the delivery of isoflurane accordingly.Begin by performing a thoracotomy to expose the heart and vessels open the chest through both sides of the rib cage. Cut from the rib edge up the axilla and transversely level with the zaphy sternum. This makes a chest wall flap.Pin the flap open to the operating board near the head. Then remove the pericardium from around the heart. Lift the inferior vena CVA with forceps and inject a milliliter of cold heparinized saline to the IVC proximal to the heart.Then attach a small artery clamp to the IVC to prevent solution from flowing back through the needle hole with gauze and a cotton applicator. Move the heart down to expose the aorta and pulmonary artery. The transverse sinus is the channel posterior to the aorta and pulmonary artery bundle.Next, cut the aorta and pulmonary artery as distally as possible to maximize the length of the vessels for the anastomosis. Now divide the IVC, the right SVC, the left SVC and the pulmonary veins. Each should be tied apart using 6.0 silk thread.First, tie the IVC and the right SVC. Then tie off the left SVC and the pulmonary veins. Now remove the heart by cutting all the vessels distal to the ties.Store the heart in cold, sterile saline at four degrees Celsius until transplantation. After preparing the animal for surgery as described for the donor, start by performing a laparotomy. Cut through the midline from the pubis to the zaphy sternum.Using paperclips bent into retractors. Open the abdomen, wrap the bowels in warm saline soaked gauze, and retract them to the upper right of the abdomen. Next, expose the infrarenal aorta and IVC between the iliac bifurcation and left renal artery and vein.Free the bundled segment of aorta and IVC. Then divide the aorta and IVC from the lumbar vessels with a calibrated cautery device. Apply small atraumatic clamps to the proximal and distal ends of the freed aorta IVC segment.Using a 30 gauge needle, punch a hole into the front wall of the aorta from the hole cut vertically making a hole the size of the donor's aorta. Take this opportunity to remove any blood clots from the aorta lumen with a rinse of heparinized saline. Now introduce the donor heart in cold saline.Position the donor heart next to the incision on the recipient's aorta. Position the donor pulmonary artery next to the recipient's IVC using 10 oh running nylon sutures. Anastomosis the aortas end to side.Start at the proximal corner and suture along the left side to the distal corner. Then rotate the animal 180 degrees. Then move the heart over and continue the sutures through the right side of the aortic wall distal to proximal.Before closing the anastomosis, gently flush the lumen with heparin eyes saline to remove clots and air. Then finish the suturing. Next, make a vertical incision in the front wall of the IVC where the pulmonary artery will be anastos.Second, use 10 oh running sutures to anastomosis. The vessels starting at the distal end of the left wall within the lumen of the IVC at the proximal end. Continue along the front right side wall and suture up to the distal end.Again.Give the lumen a good flushing before completing the anastomosis. Next place pieces of gel foam around the anastomosis. Hemostasis is achieved with gentle pressure from cotton applicators.To revascularize the recipient. Release the distal clamp first, then the proximal clamp. During the recipient operation, it's vital to optimize the accuracy and the speed of the surgery.So the cross clamp time is ideally less than 30 minute. To aid the recovery, apply warm saline at 37 degrees Celsius to the graft site. Fibrillation should be immediate and revert spontaneously to the sinus rhythm within a few minutes.Before closing the surgical site, an injection of buprenorphine should be given intramuscularly. Finish the operation by closing the wound with one five oh absorbable running suture for all the tissue layers. After closing the surgical wound, inject the recipient animal with ampicillin and a subcutaneous injection of 0.6 milliliters of saline.To help hydration place the animal on a heating pad set to 37 degrees Celsius A until it recovers consciousness. Most animals are drinking and even eating within three hours of the surgery. If after eight hours, the mouse still shows signs of distress, provide injections of buprenorphine every 12 hours until the symptoms resolve.If the mouse remains in distress for more than 48 hours, contact a veterinarian for the first 10 postoperative days. Directly monitor the grafted heartbeat by abdominal palpation. On a daily basis, record the strength of beat as four plus for a healthy graft to one plus for a weak beat due to advanced rejection.Record a non beating graft as a negative one due to complete rejection of the graft after 10 days. Continue monitoring the grafted heart pulse three times per week after 227 heterotopic heart transplants, 90.3%were successful for 24 hours, and 86.8%were successful for 48 hours. Failure included hind limb paralysis, a non beating heart due to graft, ischemic injury and or thrombosis and death graft survivors were cataloged.The cause of heart transplant failures was established by pathology. Analysis of the transplanted heart here marked by an arrow thrombosis and infarction of the heart tissue was the cause of this grafted heart's failure in the strain combination of C 57 Black six donor and recipient, no heart was rejected and the transplanted hearts survived for more than 100 days. In this case, the heart had reduced slightly in size due to muscle atrophy secondary to its non-life sustaining status.In contrast, in the rejecting strain combination of DBA two donor to C 57 Black six recipient, this transplanted heart had ceased to beat by day seven and showed evidence of rejection by 100 days. The same heart was completely fibrotic and shrunken Once marketed. This technique can be done in 90 million or less if it's performed properly, and don't forget that working with US flooring gas can be extremely hazardous.The use of an anesthetic gas scavenger is highly available.

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

Research

JoVE Journal

Medicine

The mouse heterotopic heart transplantation model has been proven by many investigators to be an important method for studying mechanisms of rejection and immune response.  However, the techniques involved are still challenging.  By modifying standard techniques we have had success with more than 1000 transplants.

Heterotopic Heart Transplantation in Mice

The murine cervical heart transplantation model is well suited for immunological as well as ischemia reperfusion injury studies. We modified the procedure using a non-suture cuff technique and performed more than 1,000 successful transplants with this approach. Herein, we provide additional details of this technique to supplement the video.

Murine Cervical Heart Transplantation Model Using a Modified Cuff Technique

The manuscript describes the steps required to perform the heterotopic heart transplant in the mouse.

Murine Heterotopic Heart Transplant Technique

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

Heterotopic Cervical Heart Transplantation in Mice

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.

Optimization of the Cuff Technique for Murine Heart Transplantation

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

Blood Circuit Reconstruction in an Abdominal Mouse Heart Transplantation Model

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

A Modified Cuff Technique for Mouse Cervical Heterotopic Heart Transplantation Model

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.

Mouse Heterotopic Cervical Cardiac Transplantation Utilizing Vascular Cuffs

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

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

This protocol presents a simplified, end-to-side suture anastomosis technique for murine cervical heart transplantation, providing a robust and reproducible model for studying transplant immunology.

Cervical Heterotopic Mouse Heart Transplantation using End-to-side Anastomosis Technique to Study Transplant Immunology

Learn a modified microsurgical technique for heterotopic heart transplantation in mice using abdominal vessel anastomosis. This method enables researchers to study transplant rejection mechanisms and strain compatibility in cardiovascular research.

A Modified Method for Heterotopic Mouse Heart Transplantion

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