Mark J. Kearns receives support from the UBC Clinician Investigator Program (awarded through the Royal College of Physicians and Surgeons of Canada), and the UBC 4YF Doctoral Fellowship.
John H. Boyd is a National Sanitorium Association and Michael Smith Foundation for Health Research Scholar. Funding through CIHR. The authors wish to thank Dr. M. Allard and Richard Wamboldt for their assistance with setup and maintenance of perfusion equipment.

All animals were housed and cared for in accordance with National and Institutional guidelines for the care and use of laboratory animals. Ethics approval for this protocol was granted by the University of British Columbia&#8217;s Animal Care Committee. Male, Sprague-Dawley rats weighing between 300 - 450 g were used for this protocol.
1. Donor Operation
<ol>
	<li>Have approximately 100 ml&#160;of cardioplegia (RT) in a cylindrical flask connected to long intravenous (IV) catheter tubing by a 3-way stopcock. Use a stand to elevate the flask to approximately 80cm above the operative surface, enabling cardioplegia delivery by gravity.</li>
	<li>To visualize structures adequately, use either a pair of surgical loupes or a dissecting microscope. 
	NOTE: We currently use a binocular operating microscope with 3.4 - 21.3X magnification.</li>
	<li>Place the donor in an anesthetic chamber and induce anesthesia with 4 - 5% isoflurane.</li>
	<li>Transfer the rat to an operating platform and maintain anesthesia by nose-cone with 1 - 2% isoflurane. Apply vet ointment to the animal&#8217;s eyes to prevent dryness. Administer midazolam (2 mg/kg) intra-peritoneally with a 25 G needle.</li>
	<li>Using surgical clippers, shave the donor from xiphisternum to mandible. Shave a small region of the left groin (for subsequent IV access). Apply a depilating agent to the operative surfaces, wait approximately 5 min, and remove hair with a piece of gauze.</li>
	<li>Prep the surgical sites with a povidone-iodine or chlorexidine-based solution (we use only chlorhexidine). Infiltrate the incisional sites with 0.1 - 0.5% lidocaine subcutaneously.</li>
	<li>Secure the forelimbs and left hindlimb to the operating platform with adhesive tape, leaving the right hindlimb free for monitoring of anesthetic depth and vital signs.</li>
	<li>After ensuring appropriate anesthetic depth by pedal pinch, make an incision in the left groin crease using a 22-blade scalpel. By blunt dissection, expose the left common femoral vein and obtain IV access as follows:
	<ol>
		<li>Gently retract the tissue overlying the femoral vein, and cannulate the vein with a 24 G IV. Connect the IV to a short length of saline-filled IV tubing, and secure the tubing in place with adhesive tape. 
		NOTE: The procedure for femoral venous cannulation in rats is described elsewhere by Jespersen and colleagues14.</li>
		<li>Connect a saline-filled 10 ml&#160;syringe to the IV tubing, and gently aspirate blood to ensure correct positioning of the IV.</li>
		<li>Inject 300 - 500 IU of unfractionated heparin through the IV tubing, and subsequently flush the tubing with 3 - 5 ml&#160;of saline.</li>
	</ol>
	</li>
	<li>Next, tracheotomize the donor as follows:
	<ol>
		<li>Make a midline incision in the soft tissues between the jugular notch and mandible using a 22-blade scalpel. Penetrate the capsule of the thyroid gland in the midline using Metzenbaum scissors, and separate its lobes using blunt dissection.</li>
		<li>Using blunt dissection, separate the strap muscles of the neck in the midline to expose the anterior surface of the trachea.</li>
		<li>Use a curved forceps to bluntly dissect a circumferential plane around the trachea. Encircle the trachea with a 4-0 silk tie.</li>
		<li>Using iris scissors, make a transverse incision in the anterior trachea, approximately 5mm inferior to the thyroid cartilage. Gently introduce the tracheal cannula (a 14 G IV) and secure it in place using the 4-0 silk tie.</li>
		<li>Connect the tracheal cannula to a mechanical ventilator. Redirect the flow of oxygen and isoflurane through the ventilator circuit, and ventilate the donor at a rate and tidal volume predicted by its weight13.</li>
	</ol>
	</li>
	<li>Make an incision in the midline of the chest (using a 22-blade scalpel), extending the incision at the jugular notch to below the xipisternum.</li>
	<li>Staying in the midline, Perform a median sternotomy using a bone cutter. Retract the edges of the sternum with a self-retaining retractor. Enter the pericardium and pleural cavities.</li>
	<li>Perform a thymectomy. It is easiest to first bluntly divide the thymus in the midline, and then separate it from surrounding structures using a combination of blunt and sharp dissection. 
	NOTE: The origin of the internal thoracic arteries may be injured when dissecting the thymus away from the superior sternal edges. To prevent bleeding, hemostatic clips may be applied before removing the thymus at these points.</li>
	<li>Using Metzenbaum scissors and/or a sharp Lauer, dissect peri-caval fat away from the inferior vena cava (IVC). Ensure that the IVC is relatively free of fat and connective tissue from the cavo-atrial junction superiorly, to the diaphragm inferiorly.</li>
	<li>Using a sharp Lauer, circumferentially free the superior vena cava (SVC) and encircle it with a 4-0 silk tie.</li>
	<li>Next, dissect the left vena cava free from surrounding structures, and ligate it proximally and distally with 4-0 silk ties. Resect the intervening portion of cava to expose the left subclavian artery.</li>
	<li>Circumferentially free the aortic arch vessels using a sharp Lauer. Apply proximal and distal surgical clips to the innominate artery, and divide it between the clips. Leave the left common carotid artery and left subclavian artery un-clipped.</li>
	<li>Next, cannulate the SVC with a 24 G IV catheter. Secure the catheter with the previously placed 4-0 silk tie.</li>
	<li>Prepare for an expedient, but gentle graft harvest.</li>
	<li>Using sharp scissors, divide the IVC just superior to the diaphragmatic surface. Divide the SVC superior to its cannulation site, ensuring that nearby airway structures aren&#8217;t injured.</li>
	<li>Turn off the ventilator and disconnect the tracheal cannula. Transect the trachea proximally.</li>
	<li>Grasp the transected trachea with forceps, and remove the heart and lungs en-bloc. This will require gentle traction with sharp dissection as the heart-lung graft is removed. Separate the trachea from the underlying esophagus. Avoid injuring the descending aorta, so that a long portion of it remains intact after harvest.</li>
</ol>
2. Preparation of the Heart-lung Graft
NOTE: while completing this portion of the procedure, an assistant should be anesthetizing the recipient rodent and preparing for graft implantation.
<ol>
	<li>Place the heart-lung graft on surgical gauze with the anterior surface of the heart facing down. Using sharp dissection, remove any residual esophagus, avoiding injury to the posterior airways.</li>
	<li>Locate the descending thoracic aorta and insert a 16 G blunt-tipped cannula. Secure the cannula in place with a 4-0 silk tie.</li>
	<li>Locate the aortic arch vessels, and apply a surgical clip to the left subclavian artery. Leave the left carotid artery un-clipped for subsequent de-airing.</li>
	<li>Administer the first dose of cardioplegia by connecting the cardioplegia tubing to the 16 G aortic cannula. The carotid artery must be temporarily occluded with forceps to ensure adequate delivery of cardioplegia. 
	NOTE: A benefit to having the aortic cannula attached is that cardioplegia may be administered continuously and/or intermittently as desired. For intermittent dosing, we typically deliver cardioplegia every 10 - 15 min in 5 ml&#160;boluses over 30 - 45 sec.</li>
	<li>Using Metzenbaum scissors and/or a sharp Lauer, dissect peri-aortic fat away from the aorta. Ensure that the aorta is relatively free of fat and connective tissue from the left subclavian artery (distal arch) to its cannulation site.</li>
	<li>Next, expose the left mainstem bronchus using a sharp Lauer, and ligate it proximally with a 4-0 silk tie. Transect the left bronchus distal to the ligature using iris scissors. Perform a left pneumonectomy by ligating the left pulmonary artery and vein with 4-0 silk ties. Transect distal to the ligature and remove the left lung.</li>
	<li>As above, remove all other lung lobes with the exception of the right upper and middle lobes. Avoid injuring the IVC in the process of performing lobectomies.</li>
	<li>Connect the cardioplegia tubing to the aortic cannula and administer continuous cardioplegia while the recipient is being prepared. Place the heart-lung graft in a sterile container (e.g., a beaker).</li>
</ol>
3. Recipient Operation
<ol>
	<li>Anesthetize the recipient as per the donor operation, above. Use vet ointment to protect the animal&#8217;s eyes from dryness. Confirm anesthesia frequently by pedal pinch.</li>
	<li>Position the animal as per the donor operation, this time leaving the right forelimb free to monitor vital signs and anesthetic depth.</li>
	<li>Shave the abdomen from xiphisternum to penis. Shave a small region of the left groin (for subsequent IV access). Apply a depilating agent to the operative surfaces, wait approximately 5 min, and remove hair with a piece of gauze.</li>
	<li>Prep the surgical sites with a povidone-iodine or chlorexidine-based solution. Infiltrate the incisional sites with 0.1 - 0.5% lidocaine subcutaneously.</li>
	<li>Tracheotomize and ventilate the recipient as directed above. Alternatively, maintain the recipient under nose-cone anesthesia.</li>
	<li>Introduce a catheter into the femoral vein as described above. Inject 300 - 500 IU of unfractionated heparin through the IV tubing, and subsequently flush the tubing with 3 - 5 ml&#160;of saline.</li>
	<li>Perform a laparotomy by making a midline abdominal incision with a 22-blade scalpel from xiphisternum to penis. Retract the abdominal wall using a self-retaining retractor. Next, retract the bowels superiorly and to the rat&#8217;s left side. Wrap them in warm, saline-soaked gauze.</li>
	<li>Expose the IVC and abdominal aorta by sharply dissecting through the overlying retro-peritoneal fat.</li>
	<li>Have a curved vessel clamp available and ready.</li>
	<li>Circumferentially free the IVC and aorta proximally and distally and encircle them with 4-0 silk ties. There should be approximately 2 - 3 cm of space between these sites.</li>
	<li>Carefully apply the curved vessel clamp, ensuring that a sufficient portion of both IVC and aorta are exposed above the jaws of the clamp.</li>
	<li>Make an incision in the anterior wall of the IVC with a 25 G needle connected to a saline-filled 1 ml&#160;syringe. Extend the incision with Potts scissors to match the length of the donor IVC orifice.</li>
	<li>Remove the heart-lung graft from its container and disconnect it from cardioplegia. Place the graft in the recipient&#8217;s abdomen in an optimal position to perform the venous anastomosis. 
	NOTE:The heart-lung graft will ultimately be oriented slightly obliquely, with the apex pointing towards the abdomen&#8217;s left lower quadrant.</li>
	<li>Secure the heel and toe ends of the anastomosis with 9-0 nylon suture. Tie a secure knot at each end, leaving the needle attached to a long arm of suture, and a short end of suture to be tied to later. Administer a dose of cardioplegia.</li>
	<li>In running fashion, complete one-half of the suture line and tie to the opposing short suture arm. Administer a dose of cardioplegia.</li>
	<li>Complete the other half of the suture line and tie it down. The venous anastomosis is complete. Administer a dose of cardioplegia</li>
	<li>Orient the heart-lung graft with the heart&#8217;s apex pointing toward the abdomen&#8217;s left lower quadrant. Ensure that the venous anastomosis is not kinked or twisted.</li>
	<li>Assess the length of donor aorta that will be required to reach the recipient aorta, maintaining both IVC and aortic anastomoses in tension-free and un-kinked configurations.</li>
	<li>Divide the descending aorta (distal to the left subclavian artery) with an iris scissors.</li>
	<li>Make an incision in the anterior wall of the recipient aorta with a 25 G needle connected to a saline-filled 1 ml&#160;syringe. Extend the incision with Potts scissors to match the length of the donor aortic orifice.</li>
	<li>Anchor the toe (superior aspect) of the donor aorta to the recipient aorta using a 9-0 nylon suture. Then, pass the needle to the medial aspect of the aorta and complete one-half of the anastomosis in running fashion.</li>
	<li>At the heel of the aortic anastomosis, reverse the orientation of the suture line (a transition in the direction of stitching), and complete the lateral half of the anastomosis in running fashion.</li>
	<li>Slowly remove the IV catheter in the SVC, and ligate the SVC with a surgical clip.</li>
	<li>De-air the aorta by locating the un-clipped left common carotid artery. Hold the artery open, in an upright position that will allow air to be freely evacuated.</li>
	<li>Briefly open the jaws of the curved vessel clamp and allow the carotid artery to bleed (de-air) for 2 - 3 sec. Re-apply the vessel clamp.</li>
	<li>Apply a surgical clip to the carotid artery. Remove the curved vessel clamp.</li>
	<li>Check for bleeding at the suture lines. If present, apply gentle compression with surgical gauze or repair with a length of 9-0 nylon suture (depending on the severity of bleeding).</li>
	<li>The donor heart should resume beating within minutes.</li>
	<li>Pay close attention to the recipient&#8217;s vital signs, depth of anesthesia, and administer volume resuscitation as needed.</li>
	<li>Depending on the experimental endpoints, either close the animal&#8217;s abdomen and let it recover from anesthesia, or prepare the animal for cardiovascular assessment. 
	NOTE: Examples of graft assessment include in vivo measures of load-dependent and load-independent hemodynamics, in vitro measures of function in Langendorff and working heart modes, and (in survival surgery) echocardiographic or MRI investigations.</li>
	<li>At the end of acute terminal experiments, animals are euthanized by exsanguination.</li>
</ol>

<ol>
	<li>Abbott, C. P., Lindsey, E. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+technique+for+heart+transplantation+in+the+rat.">A technique for heart transplantation in the rat.</a> Arch Surg. 89 (1964), 649-652 (1964).</li><li>Ma, Y., Wang, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+2+heterotopic+heart+transplant+techniques+in+rats:+cervical+and+abdominal+heart.">Comparison of 2 heterotopic heart transplant techniques in rats: cervical and abdominal heart.</a> Exp Clin Transplant. 9 (2), 128-133 (2011).</li><li>Wiedemann, D., Boesch, F., Schneeberger, S., Kocher, A., Laufer, G., Semsroth, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Graft+function+after+heterotopic+rat+heart+transplant+with+an+isolated+reperfused+working+heart:+a+methodic+consideration.">Graft function after heterotopic rat heart transplant with an isolated reperfused working heart: a methodic consideration.</a> Exp Clin Transplant. 10 (2), 154-157 (2012).</li><li>Chen, Z. H., Xia, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+technique+of+heterotopic+heart-lung+transplantation+in+the+rat.">The technique of heterotopic heart-lung transplantation in the rat.</a> J Tongji Med Univ. 6 (2), 67-70 (1986).</li><li>Ibrahim, M., Navaratnarajah, 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=Heterotopic+abdominal+heart+transplantation+in+rats+for+functional+studies+of+ventricular+unloading.">Heterotopic abdominal heart transplantation in rats for functional studies of ventricular unloading.</a> J Surg Res. 179 (1), e31-e39 (2013).</li><li>Asfour, B., Hare, J. 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=A+simple+new+model+of+physiologically+working+heterotopic+rat+heart+transplantation+provides+hemodynamic+performance+equivalent+to+that+of+an+orthotopic+heart.">A simple new model of physiologically working heterotopic rat heart transplantation provides hemodynamic performance equivalent to that of an orthotopic heart.</a> J Heart Lung Transplant. 18 (10), 927-936 (1999).</li><li>Figueiredo, J. -. L., Nahrendorf, M., Sosnovik, D. E., Weissleder, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MRI+of+a+novel+murine+working+heart+transplant+model.">MRI of a novel murine working heart transplant model.</a> Circ Heart Fail. 2 (3), 272-274 (2009).</li><li>Wen, P., Wang, X., 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=A+simple+technique+for+a+new+working+heterotopic+heart+transplantation+model+in+rats.">A simple technique for a new working heterotopic heart transplantation model in rats.</a> Transplant Proc. 45 (6), 2522-2526 (2013).</li><li>Didi&#233;, M., Biermann, 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=Preservation+of+left+ventricular+function+and+morphology+in+volume-loaded+versus+volume-unloaded+heterotopic+heart+transplants.">Preservation of left ventricular function and morphology in volume-loaded versus volume-unloaded heterotopic heart transplants.</a> A.Am J Physiol Heart Circ Physiol. 305 (4), H533-H541 (2013).</li><li>Gali&#241;anes, M., Zhai, X., Hearse, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effect+of+load+on+atrophy,+myosin+isoform+shifts+and+contractile+function:+studies+in+a+novel+rat+heart+transplant+preparation.">The effect of load on atrophy, myosin isoform shifts and contractile function: studies in a novel rat heart transplant preparation.</a> J Mol Cell Cardiol. 27 (1), 407-417 (1995).</li><li>Yokoyama, H., Ohmi, M., Murata, S., Nakame, T., Tabayashi, K., Mohri, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Proposal+of+a+working+left+heart+model+with+a+heterotopic+transplantation+technique+in+rats.">Proposal of a working left heart model with a heterotopic transplantation technique in rats.</a> J Heart Lung Transplant. 14 (4), 706-712 (1995).</li><li>Maruyama, T., Swartz, M. T., McBride, L. R., Pennington, D. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Working+heart+model+of+heterotopic+heart-lung+transplantation+in+rats.">Working heart model of heterotopic heart-lung transplantation in rats.</a> J Thorac Cardiovasc Surg. 107 (1), 210-215 (1994).</li><li>Pacher, P., Nagayama, T., Mukhopadhyay, P., B&#225;tkai, S., Kass, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+cardiac+function+using+pressure-volume+conductance+catheter+technique+in+mice+and+rats.">Measurement of cardiac function using pressure-volume conductance catheter technique in mice and rats.</a> Nat Protoc. 3 (9), 1422-1434 (2008).</li><li>Jespersen, B., Knupp, L., Northcott, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Femoral+arterial+and+venous+catheterization+for+blood+sampling,+drug+administration+and+conscious+blood+pressure+and+heart+rate+measurements.">Femoral arterial and venous catheterization for blood sampling, drug administration and conscious blood pressure and heart rate measurements.</a> J Vis Exp. (59), 3496-3410 (2012).</li><li>Habertheuer, A., Kocher, 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=Innovative,+simplified+orthotopic+lung+transplantation+in+rats).">Innovative, simplified orthotopic lung transplantation in rats).</a> J Surg Res. 185 (1), 419-425 (2013).</li></ol>

The authors have nothing to disclose.

Success with the technique described here will be predicated on several factors. Key among them will be ensuring stability of both donor and recipient animals, adopting meticulous operative technique that is safe and associated with minimal blood loss, ensuring complete cardioplegic arrest with uniform graft cooling, minimizing total ischemic time, and adequately de-airing the graft. As acknowledged above, the technique&#8217;s technical complexity is its chief limitation.
We have advanced pre...

The first rodent model of heterotopic abdominal heart transplantation (HAHT) was described by Abbott and colleagues in 19641. This technique, and subsequent modifications have been widely applied to characterize transplant graft function and immunologic status. The majority of HAHT techniques described involve a non-volume loaded heart2,3. Models of HAHT involving volume-loaded ventricles have been described, but they are frequently limited in one or more respects.
Heterotopic abdominal heart-lung transplantation (HAHLT) with a volume-loaded left ventricle (LV) has been described previously. Chen and colleagues4, and subsequently Ibrahim and colleagues5 described HAHLT with a single aorto-aortic (donor ascending to recipient abdominal aorta) anastomosis. The only volume load presented to the ventricle in this circulation is the coronary venous return. Asfour and colleagues described a HAHT technique in which the lung circuit was eliminated by anastomosing donor pulmonary artery (PA) to donor left atrium (LA)6. In this circulation, venous inflow to right ventricle (RV) occurs via a donor SVC to recipient IVC anastomosis, and the subsequent LV load is ejected into the aorto-aortic anastomosis. Cardiac function was partially assessed in vivo, and also in vitro using a Langendorff rig. Figueiredo and colleagues described a HAHLT model similar to our own7, but in mice. Venous inflow to the RV occurs via donor SVC to recipient IVC anastomosis. Blood subsequently passes through the single lung circulation and LV load is ejected into the aorto-aortic anastomosis. Cardiac function in their study was assessed by magnetic resonance imaging (MRI). Wen and colleagues described a unique HAHT technique in which the LV is loaded by means of a recipient aorta to donor LA anastomosis8. The LV, therefore, fills at systemic pressures. Cardiac function, and whether LV stroke volume is ejected antegradely in their model was not assessed.
Many of the techniques referenced above involve non-physiologic LV loading conditions, including the techniques whose partial LV load is represented only by coronary venous return. On the other hand, many techniques do approach physiologic LV loading. The majority of these techniques, as with the technique of Asfour and colleagues, omit the pulmonary circulation and utilize a donor PA to donor LA anastomosis6,9. The circulation described by Galinanes and colleagues10 employs a direct recipient cava to donor LA anastomosis, omitting the pulmonary circulation and the right heart. Yokoyama and colleagues achieve the same effect by ligating the donor PA and creating an interatrial communication in the donor heart (omitting donor lung and right heart circulations)11. The circulation of Maruyama and colleagues12 involves an anastomosis between donor left PA and recipient Ao, which permits LV filling via the pulmonary circulation as a conduit, but effectively excludes the right heart.
In cases where near physiologic loading conditions were met, we advance the technique of HAHLT in 2 major respects. First, to our knowledge, the exact configuration we report has not been described in rats. It is possibly the most versatile circulation for investigators wishing to study the physiology, structure, and immunology of the transplanted heart-lung graft. Second, we describe how the function of the transplant graft can be directly characterized in vivo. For this application, pressure-volume conductance catheters can be introduced directly into the LV apex of the transplant graft, which allows for complete cardiac functional characterization. 
The technique described here can be applied to both acute and chronic studies of transplant graft function, while the functional assessment may be performed either in vivo or in vitro. We present a model in which the loading conditions can be near physiologic, however the degree of ventricular loading may be manipulated both acutely and chronically by diverting venous return towards or away from the graft. Afterload conditions can also be manipulated. Because the lung and its airway are retained in this transplant configuration, investigators can re-ventilate the donor lung acutely. Uniquely, lung re-ventilation changes the composition of blood perfusing the transplant coronary arteries. Under non-ventilated conditions, blood ejected from the donor aorta is deoxygenated, and mixes with oxygenated blood in the recipient aorta. Under acutely ventilated conditions, ejected blood becomes oxygenated. Thus, transplant graft function can be compared under ventilated and non-ventilated conditions, and also under variably loaded conditions.
The protocol below describes important modifications to previously described HAHLT donor and recipient operations. It also describes an optimal technique for protecting the transplant graft throughout the period of ischemia (time between donor explant and recipient implant). Advantages of this technique include physiologic conditions potentially approaching that of an orthotopically transplanted graft, and a wide range of investigative applications. An important limitation is its technical complexity. With adequate mentoring and practice, the advantages of this technique will likely outweigh the challenges in adopting it.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Material/Equipment</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Celsior Cardioplegic Soution</td>
			<td>Genzyme</td>
			<td></td>
			<td>The solution is kept on ice throughout the procedure. We prepare our own solution, with slight modifications.</td>
		</tr>
		<tr>
			<td>Rodent Ventilator</td>
			<td>Harvard Apparatus</td>
			<td>Model 683</td>
			<td></td>
		</tr>
		<tr>
			<td>Vital Sign Monitor</td>
			<td>Nonin</td>
			<td>Model 9847V</td>
			<td>Displays SpO2 and heart rate.</td>
		</tr>
		<tr>
			<td>IV Cannulae</td>
			<td>Jelco</td>
			<td>3063</td>
			<td>24-26G x 3/4&#34; cannulae.</td>
		</tr>
		<tr>
			<td>IV Tubing</td>
			<td>CareFusion</td>
			<td>MP9259-C</td>
			<td>Short-length connector tubing (18cm).</td>
		</tr>
		<tr>
			<td>Surgical Clips</td>
			<td>Teleflex Medical</td>
			<td>001204</td>
			<td>Horizon titanium ligating clips.</td>
		</tr>
		<tr>
			<td>Sutures</td>
			<td>Ethicon, Sharpoint</td>
			<td>LA54G, AK-0107</td>
			<td>3-0 silk reel, and 9-0 prolene suture (single-armed, DR5 needle).</td>
		</tr>
		<tr>
			<td>Surgical Instruments</td>
			<td>Not Applicable</td>
			<td>Not Applicable</td>
			<td>The instruments used are generic, and can be purchased from any surgical supply company.</td>
		</tr>
	</tbody>
</table>

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.

The HAHLT technique described above is highly technical and requires close attention to detail. Table 1 highlights some of the key factors associated with successful versus unsuccessful procedures, and can be used as a guide for troubleshooting technical difficulties.
After the recipient aorta is unclamped, the graft coronary arteries should be seen to fill with oxygenated blood. Accordingly, the myocardium should become pink and well perfused. In technically successful experi...

Watch this Scientific Journal Video about Rat Heterotopic Abdominal Heart/Single-lung Transplantation in a Volume-loaded Configuration at JoVE.com

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

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 ...

rat-heterotopic-abdominal-heartsingle-lung-transplantation-volume

Research

JoVE Journal

Medicine

8.3K 조회수.  UBC James Hogg Research Center, St. Paul's Hospital. 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.

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

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 ...

Heterotopic Auxiliary Rat Liver Transplantation With Flow-regulated Portal Vein Arterialization in Acute Hepatic Failure

In acute hepatic failure auxiliary liver transplantation is an interesting alternative approach. The aim is to provide a temporary support until the failing native liver has regenerated.1-3 The APOLT-method, the orthotopic implantation of auxiliary segments- averts most of the technical problems. However this method necessitates extensive resections of both the native liver and the graft.4 In 1998, Erhard developed the heterotopic auxiliary liver transplantation (HALT) utilizing portal vein arterialization (PVA) (Figure 1). This technique showed promising initial clinical results.5-6 We developed a HALT-technique with flow-regulated PVA in the rat to examine the influence of flow-regulated PVA on graft morphology and function (Figure 2).
A liver graft reduced to 30 % of its original size, was heterotopically implanted in the right renal region of the recipient after explantation of the right kidney.&#160; The infra-hepatic caval vein of the graft was anastomosed with the infrahepatic caval vein of the recipient. The arterialization of the donor&#8217;s portal vein was carried out via the recipient&#8217;s right renal artery with the stent technique. The blood-flow regulation of the arterialized portal vein was achieved with the use of a stent with an internal diameter of 0.3 mm. The celiac trunk of the graft was end-to-side anastomosed with the recipient&#8217;s aorta and the bile duct was implanted into the duodenum. A subtotal resection of the native liver was performed to induce acute hepatic failure. 7
In this manner 112 transplantations were performed. The perioperative survival rate was 90% and the 6-week survival rate was 80%. Six weeks after operation, the native liver regenerated, showing an increase in weight from 2.3&#177;0.8 g to 9.8&#177;1 g. At this time, the graft&#8217;s weight decreased from 3.3&#177;0.8 g to 2.3&#177;0.8 g.
We were able to obtain promising long-term results in terms of graft morphology and function. HALT with flow-regulated PVA reliably bridges acute hepatic failure until the native liver regenerates.

Auxiliary liver transplantation provides a temporary support in acute hepatic failure, until regeneration of the failing liver. The heterotopic auxiliary liver transplantation (HALT) with portal vein arterialization (PVA) renders sufficient liver function. We developed an analogous technique in the rat, to examine the influence of the portal vein arterialization on the morphology and function of the graft.

In acute hepatic failure auxiliary liver transplantation is an interesting alternative approach. The aim is to provide a temporary support until the ...

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 ...

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 ...

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 ...

Blood Circuit Reconstruction in an Abdominal Mouse Heart Transplantation Model

The surgical technique of heterotopic abdominal heart transplantation in mice is a standard model for research in transplantation immunology. Here, the established technique for a modified blood circuit reconstruction in a heterotopic abdominal heart transplantation model is presented. This method uses the intrathoracic inferior vena cava (IIVC) instead of the pulmonary artery of the donor heart for the anastomosis to the inferior vena cava of the recipient. It is facilitating and improving success rates for abdominal heart transplantation in mice.

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

The surgical technique of heterotopic abdominal heart transplantation in mice is a standard model for research in transplantation immunology. Here, the ...

A 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.

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 ...

A Porcine Heterotopic Heart Transplantation Protocol for Delivery of Therapeutics to a Cardiac Allograft

Cardiac transplantation is the gold standard treatment for end-stage heart failure. However, it remains limited by the number of available donor hearts and complications such as primary graft dysfunction and graft rejection. The recent clinical use of an ex vivo perfusion device in cardiac transplantation introduces a unique opportunity for treating cardiac allografts with therapeutic interventions to improve function and avoid deleterious recipient responses. Establishing a translational, large-animal model for therapeutic delivery to the entire allograft is essential for testing novel therapeutic approaches in cardiac transplantation. The porcine, heterotopic heart transplantation model in the intraabdominal position serves as an excellent model for assessing the effects of novel interventions and the immunopathology of graft rejection. This model additionally offers long-term survival for the pig, given that the graft is not required to maintain the recipient's circulation. The aim of this protocol is to provide a reproducible and robust approach for achieving ex vivo delivery of a therapeutic to the entire cardiac allograft prior to transplantation and provide technical details to perform a survival heterotopic transplant of the ex vivo perfused heart.

We present a protocol for utilizing a normothermic ex vivo sanguinous perfusion system for the delivery of therapeutics to an entire cardiac allograft in a porcine heterotopic heart transplant model.

Cardiac transplantation is the gold standard treatment for end-stage heart failure. However, it remains limited by the number of available donor hearts ...

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 Mouse Model for Studying Laryngeal Transplantation

Laryngeal heterotopic transplantation, although a technically challenging procedure, offers more scientific analysis and cost benefits compared to other animal models. Although first described by Shipchandler et al. in 2009, this technique is not widely used, possibly due to the difficulties in learning the microsurgical technique and time required to master it. This paper describes the surgical steps in detail, as well as potential pitfalls to avoid, in order to encourage effective use of this technique.
In this model, the bilateral carotid arteries of the donor larynx are anastomosed to the recipient carotid artery and external jugular vein, allowing for blood flow through the graft. Blood flow can be confirmed intraoperatively by the visualization of blood filling in the graft bilateral carotid arteries, reddening of the thyroid glands of the graft, and bleeding from micro vessels in the graft. The crucial elements for success include delicate preservation of the graft vessels, making the correct size arteriotomy and venotomy, and using the appropriate number of sutures on the arterial-arterial and arterial-venous anastomoses to secure vessels without leakage and prevent occlusion. 
Anyone can become proficient in this model with sufficient training and perform the procedure in approximately 3 h. If performed successfully, this model allows for immunologic studies to be performed with ease and at low cost.

The aim of this manuscript is to describe the microsurgical steps required to perform a heterotopic laryngeal transplant in mice. The advantages of this mouse model compared to other animal models of laryngeal transplantation are its cost-effectiveness and the availability of immunologic assays and data.

Laryngeal heterotopic transplantation, although a technically challenging procedure, offers more scientific analysis and cost benefits compared to other ...