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/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>Habertheuer, A., 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=Donor+tissue-specific+exosome+profiling+enables+noninvasive+monitoring+of+acute+rejection+in+mouse+allogeneic+heart+transplantation.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Four+decades+of+vascularized+heterotopic+cardiac+transplantation+in+the+mouse.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Novel+technique+for+blood+circuit+reconstruction+in+mouse+heart+transplantation+model.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Total+arterial+anastomosis+heterotopic+heart+transplantation+model.">Total arterial anastomosis heterotopic heart transplantation model.</a> Transplantation Proceedings. 45, 625-629 (2013).</li><li>Melanie, L., et al., Moffatt-Bruce, S. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Potential+of+Heterotopic+Cardiac+Transplantation+in+Mice+as+a+Model+for+Elucidating+Mechanisms+of+Graft+Rejection.">Potential of Heterotopic Cardiac Transplantation in Mice as a Model for Elucidating Mechanisms of Graft Rejection.</a> Cardiac Transplantation. , (2012).</li><li>Westhofen, 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=The+heterotopic+heart+transplantation+in+mice+as+a+small+animal+model+to+study+mechanical+unloading+-+Establishment+of+the+procedure,+perioperative+management+and+postoperative+scoring.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Modified+Method+for+Heterotopic+Mouse+Heart+Transplantion.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Histidine-Tryptophan-Ketoglutarate+Solution+for+Donor+Heart+Preservation+Is+Safe+for+Transplantation.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modified+Suture+Technique+in+a+Mouse+Heart+Transplant+Model.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assessment+of+graft+function+in+rodent+models+of+heart+transplantation.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=cold+storage+using+a+new+histidine-tryptophan-ketoglutarate-based+preservation+solution+in+isogeneic+cardiac+mouse+grafts.">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>T&#252;rk, T. R., 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=Reduction+of+chronic+graft+injury+with+a+new+HTK-based+preservation+solution+in+a+murine+heart+transplantation+model.">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 model, the IIVC anastomosis offers a clearer sight of the operation situs enabling improved surgery of anastomoses and preventing the undesired development of too strict sutures or even damaging lesions of the vessels (Figure 4). Both the construction of the PA as well as the IIVC anastomosis are challenging even for experienced operators. The PA is very delicate and thin-walled and the IIVC is even thinner and potentially more fragile in mice. Therefore, operators need to be aware of this restriction and should apply caution when suturing the IIVC. However, at this point it is important to emphasize that although the IIVC vessel wall is very thin, it has the great advantage that the vessel length is not associated with tension to the blood vessel connection, thereby making the application of a precise suture easier and less prone to a damage. Since the cautious and safe application of the vessel sutures are very critical for a successful operation outcome, a magnification of the situs of 10 to 20-fold is recommended.
In addition, the ending of the thoracic segment of the IIVC forms a typical trumpet shaped structure (Figure 5). Its larger opening diameter represents one of the important and beneficially reasons why the IIVC can be chosen. Its usage facilitates the application of a sufficient anastomosis. This reduces both the difficulty of the operation and the operation time.
A possible event that may compromise the outcome of the transplant procedure represents a thrombosis of the vessel anastomosis, often fostered by a stenosis. Although in our model the length of the vessel does increase on purpose, this was not associated with the formation of thrombi. The trumpet shaped anastomosis of the IIVC may also exert a positive effect while greatly reducing the occurence of an anastomotic stenosis. Therefore, for this procedure a posttransplant heparinization is not necessary.
A previous analysis of the IVC-aivc&#160;anastomosis method revealed several advantages and improvements when compared to the conventional technique10. With respect to these results and from the retrospective point of view of our longstanding experience4,10,12,13, this technique results both in a reduced warm ischemia and rebeating time of the grafted heart. Even though there are no differences for longtime survival rates for this model10, the modified technique presented here is facilitating the anostomotic vessel reconstruction thus reducing the difficulty of abdominal heart transplantation in mice. The training and application of this model may therefore improve the accessibility and widespread application of abdominal heart transplantation in mice for immunological as well as cardiac research purposes.

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#160; 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&#246;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 significantly shorter than in the P-V group (27.5&#177;1.3 min, p&#60;0.01). The warm ischemia time, total recipient operation time and postoperative heart repeating time, which were 25.5&#177;1.2 min, 42.0&#177;1.5min, and 1.1&#177;0.2 min, respectively, were also significantly shortened than observed in the traditional P-V group (all p&#60;0.05) (Figure 1). Even though there are no differences for longtime survival rates in this model10,11, the modified technique facilitated abdominal heart transplantation in mice resulting both in a reduced warm ischemia and grafted heart rebeating time.
With respect to these previously published data and our experience with this model within the last 16 years, we recommend that the key operation steps should take a limited amount of time to ensure a success rate of &#62;90%4. Therefore, optimal results are achieved if the harvesting of the donor heart takes no longer than 60 min, the cold ischemia time should be limited to 40 min at maximum, and the construction of the IIVC anastomosis should not take longer than 15 min as this is directly associated with a reduced warm ischemia time.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62007/62007fig1v2.jpg" /> 
Figure 1. Comparison of procedure times between the two operation techniques (n =40 each, mean + SE) <a href="https://www.jove.com/files/ftp_upload/62007/62007fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62007/62007fig02.jpg" /> 
Figure 2. Donor heart preparation using the traditional (a, upper left) and the modified (b, lower left) mouse heart transplantation model. The photographs in panels a and b depict the aorta (A), the pulmonary artery (PA), the right atrium (RA), the left atrium (LA) and the intrathoracic inferior vena cava (IIVC) of the donor heart. Notice the difference in vessel length of the PA compared to the IIVC. Panel c shows the recipient situs prepared for the heterotopic abdominal HTX. <a href="https://www.jove.com/files/ftp_upload/62007/62007fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62007/62007fig03.jpg" /> 
Figure 3. Vascular anastomosis of the modified model: aorta (A), intrathoracic inferior vena cava (IIVC), abdominal inferior vena cava (aivc), abdominal aorta (aa). <a href="https://www.jove.com/files/ftp_upload/62007/62007fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62007/62007fig04.jpg" /> 
Figure 4. Demonstration of vessel length for the pulmonary artery (PA) and the intrathoracic inferior vena cava (IIVC). <a href="https://www.jove.com/files/ftp_upload/62007/62007fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62007/62007fig05.jpg" /> 
Figure 5. Physiological trumpet shape at the end of the intrathoracic inferior vena cava. Aorta (A), intrathoracic inferior vena cava (IIVC), right atrium (RA), left atrium (LA). The left red double arrow on the left highlights the diameter and the red circle on the right the structure of trumpet shape. <a href="https://www.jove.com/files/ftp_upload/62007/62007fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>Vena cava to vena cava (new method)</td>
			<td>Pulmonary A to vena cava(traditional method)</td>
		</tr>
		<tr>
			<td>Donor procurement (min)</td>
			<td>12.8 + 0.3</td>
			<td>11.4 + 0.4</td>
		</tr>
		<tr>
			<td>Recipient operation (min)</td>
			<td>42.0 + 1.5</td>
			<td>48.6 + 1.4**</td>
		</tr>
		<tr>
			<td>Vessel anastomosis (min)</td>
			<td>20.8 + 1.3</td>
			<td>27.5 + 1.3**</td>
		</tr>
		<tr>
			<td>Cold ischemia (min)</td>
			<td>32.8 + 0.6</td>
			<td>34.2 + 0.7</td>
		</tr>
		<tr>
			<td>Warm ischemia (min)</td>
			<td>25.5 + 1.2</td>
			<td>32.6 + 1.3**</td>
		</tr>
		<tr>
			<td>Rebeat postoperation (min)</td>
			<td>1.1 + 0.2</td>
			<td>2.1 + 0.4*</td>
		</tr>
		<tr>
			<td>Success rate</td>
			<td>92.50%</td>
			<td>90.00%</td>
		</tr>
		<tr>
			<td>*P &#60; 0.05</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>**P &#60; 0.01.</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 1. Comparison of Time Distributions and Initial Outcomes in Two Operations (n - 40, mean + SE)
Reprinted from Wu, K., Zhang, J., Fu, J., Wu, S., Philipp, T., Uwe, H., Kribben, A. and Witzke, O. Novel technique for blood circuit reconstruction in mouse heart transplantation model. Microsurgery. 26, 594-598 (2006).

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

Universidad San Sebastian

Research

JoVE Journal

Medicine

4.6K 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

Orthotopic Liver Transplantation in Rats

Clinical progress in the field of liver transplantation has been largely supported by animal models1,2. Since the publication of the first orthotopic rat liver transplantation in 1979 by Kamada et al.3, this model has remained the gold standard despite various proposed alternative techniques4. Nevertheless, its broader use is limited by its steep learning curve5.
In this video paper, we show a simple and easy-to-establish revision of Kamada's two-cuff technique. The suprahepatic vena cava anastomosis is performed manually with a running suture, and the vena porta and infrahepatic vena cava anastomoses are performed utilizing a quick-linker cuff system6. Manufacturing the quick-linker kit is shown in a separate video paper.

We present an easy-to-establish revision of the classical two-cuff technique for orthotopic liver transplantation in rat.

Clinical progress in the field of liver transplantation has been largely supported by animal models1,2. Since the publication of the first orthotopic rat ...

Using Quantitative Real-time PCR to Determine Donor Cell Engraftment in a Competitive Murine Bone Marrow Transplantation Model

Murine bone marrow transplantation models provide an important tool in measuring hematopoietic stem cell (HSC) functions and determining genes/molecules that regulate HSCs. In these transplant model systems, the function of HSCs is determined by the ability of these cells to engraft and reconstitute lethally irradiated recipient mice. Commonly, the donor cell contribution/engraftment is measured by antibodies to donor- specific cell surface proteins using flow cytometry. However, this method heavily depends on the specificity and the ability of the cell surface marker to differentiate donor-derived cells from recipient-originated cells, which may not be available for all mouse strains. Considering the various backgrounds of genetically modified mouse strains in the market, this cell surface/ flow cytometry-based method has significant limitations especially in mouse strains that lack well-defined surface markers to separate donor cells from congenic recipient cells. Here, we reported a PCR-based technique to determine donor cell engraftment/contribution in transplant recipient mice. We transplanted male donor bone marrow HSCs to lethally irradiated congenic female mice. Peripheral blood samples were collected at different time points post transplantation. Bone marrow samples were obtained at the end of the experiments. Genomic DNA was isolated and the Y chromosome specific gene, Zfy1, was amplified using quantitative Real time PCR. The engraftment of male donor-derived cells in the female recipient mice was calculated against standard curve with known percentage of male vs. female DNAs. Bcl2 was used as a reference gene to normalize the total DNA amount. Our data suggested that this approach reliably determines donor cell engraftment and provides a useful, yet simple method in measuring hematopoietic cell reconstitution in murine bone marrow transplantation models. Our method can be routinely performed in most laboratories because no costly equipment such as flow cytometry is required.

Determining donor cell engraftment presents a challenge in mouse bone marrow transplant models that lack well-defined phenotypical markers. We described a methodology to quantify male donor cell engraftment in female transplant recipient mice. This method can be used in all mouse strains for the study of HSC functions.

Murine bone marrow transplantation models provide an important tool in measuring hematopoietic stem cell (HSC) functions and determining genes/molecules ...

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

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

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

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

Rat Heterotopic Abdominal Heart/Single-lung Transplantation in a Volume-loaded Configuration

Herein, we describe a novel technique for heterotopic abdominal heart-lung transplantation (HAHLT) in rats. The configuration of the transplant graft involves anastomosis of donor inferior vena cava (IVC) to recipient IVC, and donor ascending aorta (Ao) to recipient abdominal Ao. The right upper and middle lung lobes are preserved and function as conduits for blood flow from right heart to left heart.
There are several advantages to using this technique, and it lends itself to a broad range of applications. Because the graft is transplanted in a configuration that allows for dyamic volume-loading, cardiac function may be directly assessed in vivo. The use of pressure-volume conductance catheters permits characterization of load-dependent and load-independent hemodynamic parameters. The graft may be converted to a loaded configuration by applying a clamp to the recipient&#8217;s infra-hepatic IVC. We describe modified surgical techniques for both donor and recipient operations, and an ideal myocardial protection strategy. Depending on the experimental aim, this model may be adapted for use in both acute and chronic studies of graft function, immunologic status, and variable ventricular loading conditions. The conducting airways to the transplanted lung are preserved, and allow for acute lung re-ventilation. This facilitates analysis of the effects of the mixed venous and arterial blood providing coronary perfusion to the graft.
A limitation of this model is its technical complexity. There is a significant learning curve for new operators, who should ideally be mentored in the technique. A surgical training background is advantageous for those wishing to apply this model. Despite its complexity, we aim to present the model in a clear and easily applicable format. Because of the physiologic similarity of this model to orthotopic transplantation, and its broad range of study applications, the effort invested in learning the technique is likely to be worthwhile.

We describe a novel technique for heterotopic abdominal heart-lung transplantation (HAHLT) in rats. The transplant configuration results in a partially loaded graft circulation, allowing direct functional assessment. This model may be employed for acute or chronic studies of function and immunologic status of the transplanted graft.

Herein, we describe a novel technique for heterotopic abdominal heart-lung transplantation (HAHLT) in rats. The configuration of the transplant graft ...

Apical Resection Mouse Model to Study Early Mammalian Heart Regeneration

Cardiovascular disease plagues the whole world due to intensive lifestyle changes. Heart regeneration holds great promise for repairing and restoring cardiomyocytes lost due to injury and disease. In contrast to the robust cardiac regeneration of certain lower vertebrates, adult mammalian hearts typically show minimal capacity for heart regeneration and repair. However, recent studies have sparked considerable scientific interest with the finding that, between postnatal day 1 to 7 (P1 to P7), the neonatal mouse heart retains significant regenerative potential after apical resection (i.e., surgical amputation and exposure of left ventricular apex). One major controversy over this finding might be due to the diverse surgery-related procedures used in efforts to replicate or expand upon this important finding. These instructions dynamically present the materials and methodology for apical resection in a mouse model. The salient steps of this rodent survival surgery involve hypothermia anesthesia, thoracotomy, surgical amputation of heart ventricular apex, and suture and recovery of mice. The approach described could expand the application of the apical resection mouse model for cardiovascular research.

A step-by-step video protocol of apical resection is demonstrated in this study. Apical resection is a recently highlighted surgical approach in mammalian heart regeneration research. This study may promote the application of apical resection as a standard methodology in research into the mechanism underlying heart regeneration.

Cardiovascular disease plagues the whole world due to intensive lifestyle changes. Heart regeneration holds great promise for repairing and restoring ...

Using the Sleeve Technique in a Mouse Model of Aortic Transplantation - An Instructional Video

Orthotopic aortic transplantation using the sleeve technique reduces injury to the aorta with failure rate of only 10-20%. The time to anastomose the aorta in mice using the sleeve method was short and easy averaging 20 min, permitting studies of iso/allo grafts. The following article describes the aortic transplantation procedure used in our laboratory. The mice were anesthetized with a mixture of 1.5% volume isoflurane and 100% oxygen through a face mask. At this point, the segment of the aorta between the renal arteries and its bifurcation was separated from the vena cava, freely prepared and clampedat the proximal and distal segments with a single silk suture. Prior to the removal of the aorta, a saline solution containing heparin was injected into the inferior vena cava. Then the aorta was cut between the clamps and a saline heparin solution was used to flush the lumen. The sleeve technique with monofilament sutures was used in order to transplant the abdominal aorta in the orthotopic position.

We present an orthotopic aortic transplantation model using the sleeve technique in mice. It is a very rapid anastomosis method, which can be employed in studies of vascular disease.

Orthotopic aortic transplantation using the sleeve technique reduces injury to the aorta with failure rate of only 10-20%. The time to anastomose the ...

A Chronic Cardiac Ischemia Model in Swine Using an Ameroid Constrictor

Cardiovascular disease remains the number one cause of mortality in the United States. There are numerous approaches to treating these diseases, but regardless of the approach, an in vivo model is needed to test each treatment. The pig is one of the most used large animal models for cardiovascular disease. Its heart is very similar in anatomy and function to that of a human. The ameroid placement technique creates an ischemic area of the heart, which has many useful applications in studying myocardial infarction. This model has been used for surgical research, pharmaceutical studies, imaging techniques, and cell therapies.
There are several ways of inducing an ischemic area in the heart. Each has its advantages and disadvantages, but the placement of an ameroid constrictor remains the most widely used technique. The main advantages to using the ameroid are its prevalence in existing research, its availability in various sizes to accommodate the anatomy and size of the vessel to be constricted, the surgery is a relatively simple procedure, and the post-operative monitoring is minimal, since there are no external devices to maintain. This paper provides a detailed overview of the proper technique for the placement of the ameroid constrictor.

The purpose of this protocol is to demonstrate the placement of a delayed constricting device (an ameroid constrictor) around a coronary artery in a swine model. This device creates an ischemic area of the heart that is useful for studying new diagnostic imaging techniques and new methods of treatment.

Cardiovascular disease remains the number one cause of mortality in the United States. There are numerous approaches to treating these diseases, but ...

Balloon-based Injury to Induce Myointimal Hyperplasia in the Mouse Abdominal Aorta

The use of animal models is essential for a better understanding of MH, one major cause for arterial stenosis.In this article, we demonstrate a murine balloon denudation model, which is comparable with established vessel injury models in large animals. The aorta denudation model with balloon catheters mimics the clinical setting and leads to comparable pathobiological and physiological changes. Briefly, after performing a horizontal incision in the aorta abdominalis, a balloon catheter will be inserted into the vessel, inflated, and introduced retrogradely. Inflation of the balloon will lead to intima injury and overdistension of the vessel. After removing the catheter, the aortic incision will be closed with single stiches. The model shown in this article is reproducible, easy to perform, and can be established quickly and reliably. It is especially suitable for evaluating expensive experimental therapeutic agents, which can be applied in an economical fashion. By using different knockout-mouse strains, the impact of different genes on MH development can be assessed.

This article demonstrates a murine model to study the development of myointimal hyperplasia (MH) after aortic balloon injury.

The use of animal models is essential for a better understanding of MH, one major cause for arterial stenosis.In this article, we demonstrate a murine ...

A Pre-Clinical Porcine Model of Orthotopic Heart Transplantation

Fifty-years following the first successful report, cardiac transplantation remains the gold-standard treatment for eligible patients with advanced heart failure. Multiple small-animal models of heart transplantation have been used to study the acute and long-term effects of novel therapies. However, few are tested and demonstrated success in clinical trials. It is of critical importance to evaluate new therapies in a clinically relevant large-animal model for efficient and reliable translation of basic studies&#39;&#160;findings. Here, we describe a pre-clinical large-animal (porcine) model of orthotopic heart transplantation that has been firmly established and previously used to investigate novel cardioprotective strategies. This procedure focuses on acute ischemia-reperfusion injury and is a reliable method to investigate novel interventions which have been tested and validated in smaller experimental models, such as the murine model. We demonstrate its usefulness in assessing cardiac performance during the early post-transplantation period and other potential possibilities enabled by the model.

Here, we describe a pre-clinical large-animal (porcine) model of orthotopic heart transplantation that has been firmly established and utilized to investigate novel cardioprotective strategies.

Fifty-years following the first successful report, cardiac transplantation remains the gold-standard treatment for eligible patients with advanced heart ...

A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of transplant with the potential to deliver durable heart valves capable of somatic growth with no requirement for anticoagulation. However, the immunobiology of heart valve transplants remains unexplored, highlighting the need for animal models to study this new type of transplant. Previous rat models for heterotopic aortic valve transplantation into the abdominal aorta have been described, though they are technically challenging and costly. For addressing this challenge, a renal subcapsular transplant model was developed in rodents as a practical and more straightforward method for studying heart valve transplant immunobiology. In this model, a single aortic valve leaflet is harvested and inserted into the renal subcapsular space. The kidney is easily accessible, and the transplanted tissue is securely contained in a subcapsular space that is well vascularized and can accommodate a variety of tissue sizes. Furthermore, because a single rat can provide three donor aortic leaflets and a single kidney can provide multiple sites for transplanted tissue, fewer rats are required for a given study. Here, the transplantation technique is described, providing a significant step forward in studying the transplant immunology of heart valve transplantation.

This protocol describes a simple and efficient method for the transplantation of aortic valve leaflets under the renal capsule to allow for the study of alloreactivity of heart valves.