We thank Dr. Yun Xu for her help as voice actor, Dr. med. Jianhua Peng for her help in video editing and Dr. Annika Kuckhahn for her comments and support. This work was supported in part by the German Research Foundation (DFG) to promote international collaborations (HO2581/4-1 to AH), and the National Science Foundation of China (NSFC; #81760291 to FJ).

All animal experiments were conducted following the guidelines from the directive 2010/63/EU of the European Parliament on protection of animals used for scientific purposes (Ethic committee approved, #G1071/09).
NOTE: Preliminary preparation, anesthesia, post-operative care and monitoring work are the same as performed in traditional surgical methods1,2,4. BALB/c mice served as heart donors and C57BL/6J as transplant recipients. Mice were aged 8-12 weeks, weighed~30 g at transplantation and were housed under standard conditions.
1. Preparatory steps
<ol>
	<li>For anesthesia, give mice inhalative isoflurane (2%) until they fall asleep, followed by intraperitoneal injections of ketamine (100 mg/kg) + xylazine (10 mg/kg) + acepromazine (2 mg/kg). For postoperative analgesia, apply Metamizol (200 mg/kg) p.o. and Carprofen (5 mg/kg) s.c.. 
	NOTE: The application of antibiotics was abstained on purpose as these substances may influence immunological responses.</li>
	<li>For surgery, use a set of microscopic instruments including a micro-scissor, micro-forceps, a needle holder and micro hemostatic clamps. An electrosurgical pen is also necessary. Perform sutures using 7/0er, 10/0er and&#160;4/0er nylon types.</li>
	<li>Place the mouse in a box for isoflurane inhalation (2%) for 40-60 seconds. Determine the depth of anesthesia by squeezing the paw with tweezers. If there is a complete lack of response for this stimulus, go to the next step.</li>
	<li>Once the mouse has fallen asleep, weigh the mouse.</li>
	<li>Apply an intraperitoneal injection of ketamine (100 mg/kg) + xylazine (10 mg/kg) + acepromazine (2 mg/kg) to the anesthetized mouse.</li>
	<li>Clip the abdominal fur and place the mouse on the operation table. Perform disinfection using povidone iodide for 3 times, then properly drape the mouse using a fenestrated surgical towel.</li>
</ol>
2. Donor operation procedure
<ol>
	<li>Use scissors to cut the skin from the neck to the lower abdomen, and peel off the full layer of the skin to the midline of both axillae.</li>
	<li>Use scissors to cut the muscles of the abdominal wall and gently move the viscera to the left (from the operator&#39;s view). Wrap away the viscera with a saline imbibed gauze to safely expose the inferior vena cava.</li>
	<li>Use a 1 mL syringe to inject 0.4 mL of the heparin solution (contains 500 U heparin) slowly into the lower vena cava and wait for 1 minute before pulling out the needle.</li>
	<li>Pull out the needle and use a micro-scissor to cut both the inferior vena cava and the abdominal aorta&#160;to accelerate exsanguination.</li>
	<li>Use scissors to open the chest cavity by performing a u-shaped cut; completely expose the heart, lungs and all chest blood vessels.
	<ol>
		<li>Expose the thoracic aorta, cut 1/2 of the lumen, and then cut the pulmonary vein to facilitate irrigation and drainage.</li>
		<li>Insert an irrigation tube into the opening of the thoracic aorta, inject at least 2 mL of 4 &#176;C cold histidine-tryptophan-ketoglutarate cardioplegia solution (Custodiol HTK solution)9 until the pulmonary vein outflow is completely clear and the heart completely stops beating.</li>
	</ol>
	</li>
	<li>Pull out the irrigation tube and detach the sternum.</li>
	<li>Use micro scissors to remove the thymus and to slightly strip the fat around the aortic arch.</li>
	<li>Use straight and curved forceps to expose and ligate the trunk of the arteria pulmonalis (on the right side of the aortic arch) with a 10/0 suture.</li>
	<li>Use micro forceps to separate the fat and connective tissue attached to the IIVC, expose and ligate the superior vena cava (on the left side of the aortic arch) with a 7/0 suture and use micro scissors to cut it behind the ligation.</li>
	<li>Make a 7/0 suture around the base of the heart underneath the aortic arch, the IIVC, and both auricles. Then ligate the pulmonary artery branches and venous lung vessels.</li>
	<li>Use micro scissors to transect the aortic arch as distal as possible, the lung vessels underneath the ligature and the IIVC near the diaphragm. Remove the heart out of the chest.</li>
	<li>Place the explanted donor&#39;s heart into 4 &#176;C cold HTK cardioplegia solution and preserve temporarily.</li>
</ol>
3. Recipient operation procedure
NOTE: The initial operation steps are similar to those previously shown for the donor mouse, including anesthesia and disinfection.
<ol>
	<li>Perform the abdominal skin cut in a transverse manner, cover the abdominal organs with a wet gauze using saline solution.</li>
	<li>Use micro forceps to expose the inferior vena cava and abdominal aorta and free them from surrounding fat tissue.</li>
	<li>Use micro forceps to ligate or electrocauterize side branch vessels (lateral or under the vein/aorta) below the renal vessels.</li>
	<li>Use clip applicator forceps to position two micro hemostatic clamps at the abdominal part of vein/aorta coming from the right leaving more than 1 cm distance for both aorta/vein to ensure space for the construction of the anastomosis in between them.</li>
	<li>Use micro scissors to make an incision into the aorta a little nearer to the lower clamp than to the upper clamp. Alternatively, use a 30 G needle to make a small hole and open it up with micro scissors.</li>
	<li>Position the recipient mouse so that the aorta is facing the operator with the vena cava on the other side. Then place the heart into the abdominal cavity and cover it with a small wet gauze pad.</li>
	<li>Use a 10/0 suture to adapt and stitch the donor aorta to the recipient aorta starting caudally, make a knot and proceed with a running suture to the top of the incision (about 4-5 stitches). Next, flip the heart over to the right (from the subject&#39;s view), cover it again and continue the suture on the left side until reaching the caudal end and knot it.</li>
	<li>Use an irrigation tube to inject at least 0.5 mL of 4 &#176;C HTK cardioplegia solution to flush the donor&#39;s IIVC.</li>
	<li>Use micro-scissors to cut a round hole on the abdominal inferior vena cava of the recipient, which should have the same size of the donors IIVC lumen. The incision should be located above the aortic anastomotic opening. Make the vein incision larger than the aortic incision.</li>
	<li>Use a 10/0 suture to sew the donor IIVC to the recipient vena cava starting caudally. Tie a knot and perform a running suture until the top of the incision is reached. Use five stitches and continue the suture on the left. Finally, tie a knot in the tail corner, and carefully tighten (be careful not to pull too tight).</li>
	<li>Place the small parts of the hemostatic sponge around the vein and aortic anastomoses.</li>
	<li>Use clip applicator forceps to remove first the lower and then the upper micro-hemostatic clamps and rinse the abdominal cavity with 38.0&#176;C tempered 0.9% sodium chloride.</li>
	<li>Use micro forceps to take away the hemostatic sponge.</li>
	<li>Observe the heartbeat of the transplanted heart.</li>
	<li>Use forceps to put the intestines back into the abdominal cavity and two-layer sutures (abdominal muscles followed by skin) to close the abdominal wound with a 4/0 suture.</li>
	<li>Put the mice into an oxygen and temperature control workstation chamber (e.g., INVIVO2-400) to provide a warm and oxygen rich environment for the transplanted mice to recover, wait for the mice to wake up.</li>
	<li>For postoperative analgesia, directly give Metamizol 200 mg/kg per os after operation. Four and 16 hours after operation give Metamizol 200 mg/kg per os+ Carprofen (5mg/kg) s.c. In the further followup, give Carprofen (5 mg/kg) s.c to the transplanted mice every 24 hours for three consecutive days after operation.</li>
</ol>

<ol><li>Corry, R., Winn, H., Russell, P. <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>Habertheuer, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Donor+tissue-specific+exosome+profiling+enables+noninvasive+monitoring+of+acute+rejection+in+mouse+allogeneic+heart+transplantation%5BTitle%5D)+AND+%22Journal+of+Thoracic+and+Cardiovascular+Surgery%22%5BJournal%5D)">Donor tissue-specific exosome profiling enables noninvasive monitoring of acute rejection in mouse allogeneic heart transplantation.</a> Journal of Thoracic and Cardiovascular Surgery. 155 (6), 2479-2489 (2018).</li>
<li>Plenter, R. J., Zamora, M. R., Grazia, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Four+decades+of+vascularized+heterotopic+cardiac+transplantation+in+the+mouse%5BTitle%5D)+AND+%22Journal+of+Investigative+Surgery%22%5BJournal%5D)">Four decades of vascularized heterotopic cardiac transplantation in the mouse.</a> Journal of Investigative Surgery. 26, 223-228 (2013).</li>
<li>Wu, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Novel+technique+for+blood+circuit+reconstruction+in+mouse+heart+transplantation+model%5BTitle%5D)+AND+%22Microsurgery%22%5BJournal%5D)">Novel technique for blood circuit reconstruction in mouse heart transplantation model.</a> Microsurgery. 26, 594-598 (2006).</li>
<li>Alamran, F. G., Shahkolahi, M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Total+arterial+anastomosis+heterotopic+heart+transplantation+model%5BTitle%5D)+AND+%22Transplantation+Proceedings%22%5BJournal%5D)">Total arterial anastomosis heterotopic heart transplantation model.</a> Transplantation Proceedings. 45, 625-629 (2013).</li>
<li>Melanie, L., et al. Potential of Heterotopic Cardiac Transplantation in Mice as a Model for Elucidating Mechanisms of Graft Rejection. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aMelanie%2C+L+%22Cardiac+Transplantation%22">Cardiac Transplantation</a>. Moffatt-Bruce, S. D., et al. , IntechOpen (2012).</li>
<li>Westhofen, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+heterotopic+heart+transplantation+in+mice+as+a+small+animal+model+to+study+mechanical+unloading+-+Establishment+of+the+procedure%2C+perioperative+management+and+postoperative+scoring%5BTitle%5D)+AND+%22PLoS+One%22%5BJournal%5D)">The heterotopic heart transplantation in mice as a small animal model to study mechanical unloading - Establishment of the procedure, perioperative management and postoperative scoring.</a> PLoS One. 14 (4), 0214513(2019).</li>
<li>Wang, C., Wang, Z., Allen, R., Bishop, G. A., Sharland, A. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+Modified+Method+for+Heterotopic+Mouse+Heart+Transplantion%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%22%5BJournal%5D)">A Modified Method for Heterotopic Mouse Heart Transplantion.</a> Journal of Visualized Experiments. (88), e51423(2014).</li>
<li>Mokbel, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Histidine-Tryptophan-Ketoglutarate+Solution+for+Donor+Heart+Preservation+Is+Safe+for+Transplantation%5BTitle%5D)+AND+%22Annals+of+Thoracic+Surgery%22%5BJournal%5D)">Histidine-Tryptophan-Ketoglutarate Solution for Donor Heart Preservation Is Safe for Transplantation.</a> Annals of Thoracic Surgery. 109 (3), 763-770 (2020).</li>
<li>Song, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Modified+Suture+Technique+in+a+Mouse+Heart+Transplant+Model%5BTitle%5D)+AND+%22Asian+Journal+of+Surgery%22%5BJournal%5D)">Modified Suture Technique in a Mouse Heart Transplant Model.</a> Asian Journal of Surgery. 34 (2), 86-91 (2011).</li>
<li>Martins, P. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Assessment+of+graft+function+in+rodent+models+of+heart+transplantation%5BTitle%5D)+AND+%22Microsurgery%22%5BJournal%5D)">Assessment of graft function in rodent models of heart transplantation.</a> Microsurgery. 28 (7), 565-570 (2008).</li>
<li>Wu, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((cold+storage+using+a+new+histidine-tryptophan-ketoglutarate-based+preservation+solution+in+isogeneic+cardiac+mouse+grafts%5BTitle%5D)+AND+%22European+Heart+Journal%22%5BJournal%5D)">cold storage using a new histidine-tryptophan-ketoglutarate-based preservation solution in isogeneic cardiac mouse grafts.</a> European Heart Journal. 32 (4), 509-516 (2011).</li>
<li>Türk, T. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Reduction+of+chronic+graft+injury+with+a+new+HTK-based+preservation+solution+in+a+murine+heart+transplantation+model%5BTitle%5D)+AND+%22Cryobiology%22%5BJournal%5D)">Reduction of chronic graft injury with a new HTK-based preservation solution in a murine heart transplantation model.</a> Cryobiology. 64 (3), 273-278 (2012).</li>
</ol>

The surgical technique of heterotopic abdominal heart transplantation in mice is very challenging and this implicates a restriction to the widespread use of this model.
One of the disadvantages of the conventional technique is the limiting length of the donor&#39;s pulmonary artery (PA). It is usually of about 2 mm of length, whereas the length of the IIVC of the donor heart used in our model is generally about 1 cm (Figure 2). That means that in the modified mode...

The surgical technique of heterotopic abdominal heart transplantation in mice represents a standard model for research in transplantation immunology1,2,3. However, it is very challenging to perform and this implicates a restriction to the widespread use of this model4,5.
In traditional mouse heart transplantation (THTx), the donor aorta and the recipient abdominal aorta are anastomosed while the pulmonary artery is anastomosed to the recipient inferior vena cava6,7,8.
In this modified mouse heart transplantation technique, the donor aorta is anastomosed to the recipient abdominal aorta and the donor IIVC is anastomosed to the recipient inferior vena cava(3,4,6) (Figure 2 and Figure 3).

<table><tbody><tr><td>30G-needles</td><td>Braun</td><td>456300</td><td></td></tr><tr><td>acepromazine</td><td>CP Pharma</td><td>Tranquisol P</td><td></td></tr><tr><td>BALB/c AnNCrl mice</td><td>Charles River. Germany</td><td>no catalog number</td><td></td></tr><tr><td>Bepanthen eye ointment</td><td>Haus-Apotheke</td><td>PZN 01578675</td><td></td></tr><tr><td>Bonn Micro Forceps</td><td>FST</td><td>11083-07</td><td></td></tr><tr><td>Box for insulation and oxygen supply device</td><td>RUSKINN</td><td>INVIV</td><td></td></tr><tr><td>C57BL/6J&amp;nbsp; mice</td><td>Charles River. Germany</td><td>no catalog number</td><td></td></tr><tr><td>Carprofen</td><td>Zoetis</td><td>Rimadyl 50 mg/ml</td><td></td></tr><tr><td>CATHETER-FEP 26G</td><td>TERUMO</td><td>Surflo-W</td><td></td></tr><tr><td>Clip Applicator Forceps Style</td><td>FST</td><td>18057-14</td><td></td></tr><tr><td>Curved forceps</td><td>WPI</td><td>14114-G</td><td></td></tr><tr><td>custodiol/HTK</td><td>Dr. Franz K&amp;ouml;hler Chemie</td><td>no catalog numer</td><td></td></tr><tr><td>Cutasept skin disinfection</td><td>VWR</td><td>BODL980365</td><td></td></tr><tr><td>electrosurgical pen</td><td>Bovie</td><td>CHANGE-A-TIP</td><td></td></tr><tr><td>gauze pads, cotton swabs</td><td>Lohmann-Rauscher</td><td>13353</td><td></td></tr><tr><td>Heating mat</td><td>THERMO MAT PRO 30W</td><td>HTP-30</td><td></td></tr><tr><td>Hemostatic sponge</td><td>CuraSpon</td><td>J1276A</td><td></td></tr><tr><td>heparine-solution</td><td>Haus-Apotheke</td><td>PZN 03029820</td><td></td></tr><tr><td>Ice box</td><td>PETZ</td><td>No Catalog Number available</td><td></td></tr><tr><td>Inhalation anesthesia device</td><td>GROPPLER</td><td>BKGM 0616</td><td></td></tr><tr><td>insulation and oxygen supply device</td><td>RUSKINN</td><td>INVIV</td><td></td></tr><tr><td>isoflurane</td><td>CP Pharma</td><td>Isofluran CP 1 ml/ml</td><td></td></tr><tr><td>ketamine</td><td>Zoetis</td><td>no catalog numer</td><td></td></tr><tr><td>metamizole</td><td>WDT</td><td>no catalog numer</td><td></td></tr><tr><td>Micro scissors</td><td>FST</td><td>15000-00,15000-10</td><td></td></tr><tr><td>Micro Serrefine ( Clamp ) Angled / 16 mm</td><td>FST</td><td>18055-06</td><td></td></tr><tr><td>Microscope</td><td>Leica</td><td>LEICAMZ6</td><td></td></tr><tr><td>Microscope light</td><td>SCHOTT</td><td>KL2500LED</td><td></td></tr><tr><td>Saline solution (NaCl 0.9%)</td><td>Haus-Apotheke</td><td>PZN 06178437</td><td></td></tr><tr><td>Scissors</td><td>Peha Instruments</td><td>991083/4</td><td></td></tr><tr><td>small Petri dish</td><td>Sarstedt</td><td>833900</td><td></td></tr><tr><td>Straight forceps</td><td>WPI</td><td>14113-G</td><td></td></tr><tr><td>surgical tape</td><td>BSN</td><td>4120</td><td></td></tr><tr><td>Suture Tying Forceps - 10 cm</td><td>FST</td><td>18025-10</td><td></td></tr><tr><td>Sutures(10-0)</td><td>Medtronic</td><td>N2540</td><td></td></tr><tr><td>Sutures(4-0)</td><td>ETHILON</td><td>V4940H</td><td></td></tr><tr><td>Sutures(7-0)</td><td>ETHILON</td><td>1647H</td><td></td></tr><tr><td>Syringe (0.3 mL)</td><td>BD</td><td>324826</td><td></td></tr><tr><td>Syringe (1 mL)</td><td>BD</td><td>320801</td><td></td></tr><tr><td>xylazine</td><td>Bayer</td><td>Rompun</td><td></td></tr></tbody></table>

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.

Here, a modified technique of heterotopic abdominal heart transplantation in mice that has been previously developed in our laboratory and has proven useful for the last 16 years is presented. Previously, it was reported that in overall 40 cases of vena cava to vena cava (V-V group) compared to 40 cases of the traditional pulmonary artery to vena cava (P-V group) anastomosis procedure4 (Table 1) the vessel anastomosis&#160;took 20.8&#177;1.3 min&#160;in the V-V group, which was si...

Watch this Scientific Journal Video about Blood Circuit Reconstruction in an Abdominal Mouse Heart Transplantation Model at JoVE.com

Blood Circuit Reconstruction in an Abdominal Mouse Heart Transplantation Model

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 ...

blood-circuit-reconstruction-an-abdominal-mouse-heart-transplantation

Research

JoVE Journal

Medicine

4.7K Views.  University Hospital Erlangen, Friedrich-Alexander University Erlangen-Nürnberg. A novel technique for blood circuit reconstruction in a heterotopic abdominal mouse heart transplantation model is demonstrated.

Video: Blood Circuit Reconstruction in an Abdominal Mouse Heart Transplantation Model

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 ...

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

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 Cuff Technique for Mouse Cervical Heterotopic Heart Transplantation Model

Cardiac allograft rejection limits the long-term survival of patients after heart transplantation. A mouse heart transplantation model is ideal for investigating the mechanism of cardiac allograft rejection in preclinical studies because of their high homology with human genes. This understanding would help develop unique approaches to improving patients&#39; long-term survival treated with cardiac allografts. In a mouse model, abdominal donor heart implantation is commonly performed with an end-to-end&#160;anastomosis to the recipient&#39;s aorta and inferior vena cava using stitches. In this model, the donor&#39;s heart is implanted by end-to-end anastomosis to the recipient&#39;s carotid artery and jugular vein by the modified-Cuff technique. The transplantation surgery is performed without stitching and thus may increase the survival of the recipient since there is no interference with the blood supply and venous reflux of the lower body. This mouse model would help investigate the mechanisms underlying the immunological and pathological (acute/chronic) rejection of cardiac allografts.

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

Cardiac allograft rejection limits the long-term survival of patients after heart transplantation. A mouse heart transplantation model is ideal for ...

A Heterotopic Rat Heart Transplantation Model using Circulatory Death Donor Hearts

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

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

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