The authors have no acknowledgements.

The institutional animal care committee approved all experimental protocols and animals were treated following the &#8220;Guide for the Care and Use of Laboratory Animals&#8221; prepared by the Institute of Laboratory Animal Resources, National Research Council, 1996. Male Yorkshire pigs (40&#8211;50 kg) were used to perform the orthotopic heart transplants (animal size can vary according to the investigators discretion and experimental goals).
1. Donor Procedure
<ol>
	<li>Anesthetic induction and animal preparation:
	<ol>
		<li>Premedicate the animal using an intramuscular injection of Ketamine (20 mg/kg), Midazolam (0.3 mg/kg), and Atropine (0.04 mg/kg). Perform anesthetic induction and maintenance using inhalational Isoflurane (end tidal concentration: 1%&#8211;3%) + 3 L/min O2 via face mask.</li>
		<li>Confirm anesthetic adequacy by ensuring a relaxed jaw tone and an absence of pain during toe pinch. Anesthetic adequacy needs to follow institutional guidelines.</li>
		<li>Once anesthetic adequacy is confirmed, perform an orotracheal intubation using an endotracheal tube of size 6.5&#8211;8 mm.</li>
		<li>Place an oxygen saturation monitor on the ear or bottom lip for continuous monitoring. Place the cautery electrode pad on the animal&#8217;s back.</li>
		<li>Insert a peripheral intravenous access via ear vein (e.g. 20 g Angiocath). Start a maintenance infusion (e.g. 0.9% NaCl). Administer 2 g of Magnesium Sulfate to prevent arrhythmias.</li>
		<li>Insert a percutaneous central venous sheath introducer using the Seldinger technique into the right jugular vein if right-sided heart catheterization and cardiac output measurements are to be performed. Place the animal in Trendelenburg position to facilitate venous access. Alternatively, this can be done in the left internal jugular vein. If access cannot be established, this step may be performed following midline sternotomy by dissecting the internal jugular vein (left or right) and inserting the sheath directly.</li>
	</ol>
	</li>
	<li>Donor heart procurement:
	<ol>
		<li>Perform a midline sternotomy from the mid cervical region to below the xiphoid process using a cautery pen. Open the sternum with a bone saw. Ensure adequate hemostasis throughout the entire procedure (e.g. cauterize sternum, and/or application of bone wax).</li>
		<li>In the cervical region, retract the sternocleidomastoid muscle to the center and dissect the right carotid artery (alternatively, this can be done with the left carotid artery). Place an arterial access line (e.g. 20 g Angiocath) into the right carotid artery for invasive arterial pressure monitoring.</li>
		<li>Dissect and remove the thymus from over the pericardium. Gently lift the thymus from the pericardium, using the cautery, and dissect the structure from the pericardium. To prevent bleeding, cauterize the small vessels that originate from the aorta and superior vena cava (SVC) to irrigate the thymus.</li>
		<li>Open the pericardium. Dissect the aortopulmonary space using the cautery. For this, have the assistant retract the right ventricular outflow track inferiorly and the pulmonary artery to the left and have the surgeon retract the aorta to the right. Do this carefully to avoid direct lesions to the pulmonary artery.</li>
		<li>Clear the anterior aspect of the ascending aorta from connective tissue. Carefully retract the right ventricular outflow tract and place a purse-string suture using a 4-0 prolene on the proximal ascending aorta adventitia (avoid full thickness sutures). Secure this suture with a tourniquet.</li>
		<li>Administer 30,000 units of heparin (&#62;300 U/Kg) to achieve systemic anticoagulation. Insert the cardioplegia delivery cannula (e.g. DLP aortic root cannula) into the ascending aorta between the purse-string suture previously placed and secure by tightening the tourniquet. Prepare the cardioplegic solution that will be used and connect to the delivery cannula.</li>
		<li>Open the SVC and inferior vena cava (IVC), and the left inferior pulmonary vein to ensure adequate cardiac venting. Place the aortic cross-clamp on the distal ascending aorta (above the cardioplegia cannula). Alternatively, the left ventricle can be vented by opening the left atrial appendage and placing suction.</li>
		<li>Initiate cardioplegia infusion targeting an aortic root pressure of 80 to 100 mmHg. 
		NOTE: The authors applied a model using 1.5 L of a standard extracellular hyperkalemic cardioplegic solution at 4 &#176;C. Different solutions and volumes can be used according to the experimental set-up.</li>
		<li>Place ice slush (0.9% NaCl) in the thoracic cavity and over the organ for cooling following initiation of cardioplegia. After the cardioplegia infusion is finished, proceed with cardiectomy in a traditional manner. Section the aorta and the pulmonary artery after the innominate artery and at the bifurcation, respectively, to ensure sufficient length for implant.</li>
		<li>After removal, place the organ in an organ bag with at least 500 mL of preservation solution (standard extracellular hyperkalemic solution). Place this on ice and keep at 4 &#176;C. This step can be modified according to the experimental design and objective.</li>
	</ol>
	</li>
</ol>
2. Recipient Procedure:
<ol>
	<li>Anesthetic induction and animal preparation:
	<ol>
		<li>Perform anesthesia and monitoring as described in the donor procedure (steps 1.1.1 to 1.1.6).</li>
		<li>After inserting the percutaneous central venous sheath introducer into the right jugular vein, insert a central venous catheter (e.g. double-lumen) into the left jugular vein using the Seldinger technique. Alternatively, this can be done after the midline sternotomy as described above.</li>
	</ol>
	</li>
	<li>Cardiopulmonary bypass (CPB):
	<ol>
		<li>Perform the midline sternotomy and expose the heart and great vessels as described in the donor procedure (steps 1.2.1 to 1.2.4).</li>
		<li>Dissect between the SVC and the innominate artery, and the IVC and pericardium using the Metzenbaum and &#8220;right-angle&#8221; forceps. Encircle the SVC and IVC using an umbilical tape (alternatively a simple O silk suture can be used). Secure each tape/suture with a tourniquet.</li>
		<li>Place 2 concentric purse-string sutures using a 4-0 prolene suture on the distal ascending aorta adventitia (avoid full thickness sutures). Place purse-string sutures using 4-0 prolene on the IVC and the SVC at the level of the pericardial reflection. Secure these sutures with tourniquets.</li>
		<li>During CPB preparation, have an assistant setup and prime the system according to the investigators&#8217; and experimental needs. The current procedure uses the same setup used as the institution&#8217;s Cardiovascular Surgery Division and employs the help of a trained perfusionist. The bypass system is primed with 2 litres of crystalloid solution (e.g. Plasmalyte) with 500 mg of Solumedrol.</li>
		<li>Administer 30,000 units of heparin (&#62;300 U/Kg) to achieve systemic anticoagulation. Activated clotting time (ACT) should be above 300 s, if the test is available.</li>
		<li>Cannulate the aorta with a 17 to 21 F arterial cannula. Use an extracorporeal membrane oxygenator (ECMO) cannula (e.g. EOPA arterial cannula) inserted using the Seldinger technique to facilitate this step and avoid blood loss. Alternatively, a standard bypass arterial cannula can be used.</li>
		<li>Connect the cannula to the arterial line of the bypass circuit using a 3/8-3/8 connector. Ensure complete deairing to avoid air embolism.</li>
		<li>Perform a bicaval cannulation. For this, cannulate the SVC and then the IVC using 24 to 28 F right angled single-stage venous cannula (e.g. DLP single lumen angled venous cannula).
		<ol>
			<li>First, make a small incision (5 mm) at the center of the purse-string suture. Dilate the incision with a small angled instrument (e.g. right angle or a snap). Insert the cannula directing the angle superiorly in the SVC and inferiorly in the IVC (away from the heart). Secure by tightening the tourniquet holding the purse-string suture.</li>
			<li>Between each step, cover the incision with a finger to avoid excessive bleeding. Connect the cannulas to the venous line of the bypass circuit using a 3/8-3/8-1/2 Y connector. Ensure deairing to avoid an airlock in the system.</li>
		</ol>
		</li>
		<li>Initiate CPB. Adjust flowrates to maintain an arterial pressure above 50 mmHg (approximately 4 L/min). Maintain normothermia throughout the procedure. 
		&#8203;NOTE: These settings can be modified according to the experimental design. Vasoactive medications should only be administered during CPB if needed to aid in pressure regulation (e.g. epinephrine infusion).</li>
	</ol>
	</li>
	<li>Donor heart implantation:
	<ol>
		<li>After initiation of CPB, open the left pleura and retract the native heart to the right. Dissect and encircle the left hemiazygous vein using a sharp dissection instrument (e.g. Metzenbaum) and a right-angle, respectively. Ligate distally with an O Silk tie. Only one ligature is needed as the native heart will be removed.</li>
		<li>Cross-clamp the recipient aorta proximally to the arterial cannula. Snare both vena cava with the tourniquets previously placed using O silk ties. Remove the recipient&#8217;s native heart. Alternatively, the heart can be arrested using standard hypothermic blood-based cardioplegia.</li>
		<li>During cardiectomy, make sure to maintain large cuffs in the recipient to facilitate donor heart implant. For this, section the aorta and pulmonary artery proximally, close to their roots. Similarly, the left and right atriums must be kept with large cuffs. Leave the right and left atrial appendages in the recipient cuffs, which may be needed during the anastomoses.</li>
		<li>Prepare the donor heart for implant.
		<ol>
			<li>Dissect the pulmonary artery off the left atrium and separate the pulmonary artery completely from the aorta. Leave at least 2-3 cm of each vessel to be trimmed during implant as needed. Ligate both vena cava with an O silk tie. Unite all pulmonary veins, creating a single left atrial cuff to be anastomosed.</li>
			<li>Compare left atrial cuff sizes (donor and recipient) and trim each as needed to become similar sizes. The recipient&#8217;s left atrial appendage can be shortened, or the donor&#8217;s left atrial roof and appendage can be opened for this.</li>
		</ol>
		</li>
		<li>Deliver the first cardioplegia dose to the donor heart using the previously placed cardioplegia cannula as described in step 1.2.6. Cardioplegic protective solution consists of 500 mL of a 2:1 mixture of blood:crystalloid containing 24 mEq of potassium and delivered at 10&#176;C. Achieve the desired potassium concentration by adding potassium chloride to the cardioplegic mixture.</li>
		<li>After the completion of each anastomosis, deliver an additional dose of 300 mL of cardioplegia at 10 &#176;C containing 8 mEq of potassium.</li>
		<li>Following all anastomoses and before removal of the aortic cross-clamp, administer an additional dose of 500 mL of warm (37 &#176;C) blood cardioplegia with 8 mEq of potassium.</li>
		<li>Remove the donor heart from storage and implant with the standard biatrial anastomotic technique in the following sequence: left atrium, right atrium, pulmonary artery, and aorta. Use a 4-0 prolene suture with a SH needle for the left and right atrium, and the aorta and a 5-0 prolene suture with a BB needle for the pulmonary artery.
		<ol>
			<li>Left atrium: place a 4-0 prolene suture at the junction between the left atrium and the IVC (right inferior margin) and another at 180&#176; from the first, connecting the donor and recipient cuffs. Complete the posterior wall anastomosis. Complete the anterior wall anastomosis. This is performed from the superior suture to the inferior suture.</li>
			<li>Right atrium: open the donor right atrium from the appendage towards the IVC, creating a donor cuff that matches the recipient cuff size. Starting at the inferior angle (junction between IVC and right atrium), complete the interior wall anastomosis and then the lateral wall.</li>
			<li>Pulmonary artery: trim the edges of both recipient and donor pulmonary arteries to create matching sizes. Place a 5-0 prolene suture at the left lateral wall uniting donor and recipient vessels and another at the right lateral edge. Complete the inferior wall anastomosis and then the anterior wall anastomosis.</li>
			<li>Aorta: trim as described with the pulmonary artery. Place a suture on the left lateral wall connecting donor and recipient vessels. Complete the inferior wall and then the anterior wall anastomosis.</li>
			<li>Perform single layered anastomoses, except for the pulmonary artery, where a double layered anastomosis is required. The porcine tissue is extremely fragile and should be handled carefully to avoid tears. Importantly, the pulmonary artery anastomosis is the most delicate step of the implant and must be done with extreme care. The implant technique can be modified according to the surgeons&#8217; preference and the experimental design.</li>
		</ol>
		</li>
		<li>Following completion of all anastomoses and warm cardioplegia dose delivery as described above, remove the aortic cross-clamp. Inspect all anastomoses for sites of bleeding, they should be corrected at this point.</li>
		<li>Reperfuse the donor heart for 60 min on CPB. Ventricular arrhythmias can be treated with internal defibrillation (20-50 J). If required, ventricular pacing can be used to maintain a heart rate of 100 beats per min. Antiarrhythmic drugs (e.g. Amiodarone, Lidocaine, or Magnesium Sulphate) can be used at the investigators discretion, if required.</li>
		<li>After 60 min of reperfusion, administer 1 g of calcium chloride. Initiate weaning from CPB by decreasing the flow to half, then one-quarter and then off. The central venous line can be used to monitor central venous pressure, targeting 10 mmHg. Initiate infusion of vasoactive and inotropic medications (e.g. dobutamine, epinephrine, norepinephrine, and vasopressin) according to the experimental design or the investigators&#8217; discretion.</li>
		<li>Weaning is deemed successful if the animal maintains a systolic arterial pressure above 60 mmHg for over 30 min after discontinuation of CPB. As this is not a survival model, do not reverse the heparin; continuous bleeding can occur from needle holes and dissected structures (e.g sternum). Donor hearts respond well to small and repeated doses of volume replacement using the CPB system. Additionally, the porcine model responds well to dobutamine. 
		NOTE: The recipient management should be tailored to the investigators&#8217; experience and the experimental design. A cardiac anesthesiologist may aid in this manner.</li>
	</ol>
	</li>
</ol>
3. Graft Assessment:
<ol>
	<li>Functional assessment:
	<ol>
		<li>This large animal model has the advantage of having an open chest approach at all times, which facilitates direct functional assessment. To measure cardiac contractility, use pressure-volume (PV) loop analyses, echocardiography, and/or right-sided catheterization.</li>
		<li>Pressure-volume loops10: Place an umbilical tape around the IVC, and insert a PV conductance catheter into the left ventricle through a small apical ventriculotomy to permit continuous measurements of left ventricular PV relations. Obtain steady-state recordings to generate volume-dependent parameters (e.g. developed pressure and stroke work) and then obtain occlusion recordings in triplicate by occluding the IVC to generate volume-independent parameters (e.g. preload recruitable stroke work).</li>
		<li>Echocardiography: have cardiac anesthesiologists obtain epicardial images using a standard transesophageal probe.</li>
		<li>Right-sided catheterization: Insert a Swans-Ganz catheter through the venous sheath placed at the beginning of the procedure and guid towards the pulmonary artery. This enables the measurement of central venous pressure, right ventricular pressure, pulmonary artery pressures, pulmonary capillary wedge pressure, and cardiac output using the thermodilution technique.</li>
		<li>Perform contractile evaluation at baseline and following 2 and 3 h post-reperfusion of the donor heart in the recipient. This can be modified by the investigators according the experimental design.</li>
	</ol>
	</li>
	<li>Metabolic assessment:
	<ol>
		<li>For metabolic assessments, collect arterial and venous (alternatively: mixed venous) blood samples and store the plasma for subsequent analyses. Real-time blood gas analyses and lactate levels should also be obtained.</li>
		<li>Collect these samples at baseline in the donor, before procurement in the donor, at baseline in the recipient, and at 15, 30 and 60 min of reperfusion of the donor heart (after removing cross-clamp). This can be modified according to the experimental design.</li>
	</ol>
	</li>
	<li>Experiment termination and euthanasia:
	<ol>
		<li>Once all assessments are finished, exsanguinate the recipient animal into the venous reservoir of the CPB circuit by opening the venous line clamp. Alternatively, exsangination can be achieved by harvesting the cardiac alograft to collect samples (i.e. myocardial biopsies).</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Lund, L. H., Edwards, L. B., 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+Registry+of+the+International+Society+for+Heart+and+Lung+Transplantation:+Thirty-third+Adult+Heart+Transplantation+Report-2016;+Focus+Theme:+Primary+Diagnostic+Indications+for+Transplant.">The Registry of the International Society for Heart and Lung Transplantation: Thirty-third Adult Heart Transplantation Report-2016; Focus Theme: Primary Diagnostic Indications for Transplant.</a> The Journal of Heart and Lung Transplantation. 35 (10), 1158-1169 (2016).</li><li>Lund, L. H., Edwards, L. B., 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+Registry+of+the+International+Society+for+Heart+and+Lung+Transplantation_+Thirty-first+Official+Adult+Heart+Transplant+Report-2014;+Focus+Theme_+Retransplantation.">The Registry of the International Society for Heart and Lung Transplantation_ Thirty-first Official Adult Heart Transplant Report-2014; Focus Theme_ Retransplantation.</a> The. Journal of Heart and Lung Transplantation. 33 (10), 996-1008 (2014).</li><li>Cos&#237;o Carmena, M. D. G., G&#243;mez Bueno, 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=Primary+graft+failure+after+heart+transplantation:+characteristics+in+a+contemporary+cohort+and+performance+of+the+RADIAL+risk+score.">Primary graft failure after heart transplantation: characteristics in a contemporary cohort and performance of the RADIAL risk score.</a> The Journal of Heart and Lung Transplantation. 32 (12), 1187-1195 (2013).</li><li>Fedak, P. W. M., Rao, V., 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=Combined+endothelial+and+myocardial+protection+by+endothelin+antagonism+enhances+transplant+allograft+preservation.">Combined endothelial and myocardial protection by endothelin antagonism enhances transplant allograft preservation.</a> The Journal of Thoracic and Cardiovascular Surgery. 129 (2), 407-415 (2005).</li><li>Ramzy, D., Rao, V., 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=Cardiac+allograft+preservation+using+donor-shed+blood+supplemented+with+L-arginine.">Cardiac allograft preservation using donor-shed blood supplemented with L-arginine.</a> The Journal of Heart and Lung Transplantation. 24 (10), 1665-1672 (2005).</li><li>Rao, V., Feindel, C. M., Weisel, R. D., Boylen, P., Cohen, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Donor+blood+perfusion+improves+myocardial+recovery+after+heart+transplantation.">Donor blood perfusion improves myocardial recovery after heart transplantation.</a> Journal of Heart and Lung Transplantation. 16 (6), 667-673 (1997).</li><li>Wicomb, W. N., Cooper, D. K., Barnard, C. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Twenty-four-hour+preservation+of+the+pig+heart+by+a+portable+hypothermic+perfusion+system.">Twenty-four-hour preservation of the pig heart by a portable hypothermic perfusion system.</a> Transplantation. 34 (5), 246-250 (1982).</li><li>Badiwala, M. V., Ramzy, 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=Donor+pretreatment+with+hypertonic+saline+attenuates+primary+allograft+dysfunction:+a+pilot+study+in+a+porcine+model.">Donor pretreatment with hypertonic saline attenuates primary allograft dysfunction: a pilot study in a porcine model.</a> Circulation. 120, 206-214 (2009).</li><li>Ribeiro, R. V. P., Badiwala, M. V., Ramzy, D., Tumiati, L. C., Rao, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recipient+Hypertonic+Saline+Infusion+Prevents+Cardiac+Allograft+Dysfunction.">Recipient Hypertonic Saline Infusion Prevents Cardiac Allograft Dysfunction.</a> The Journal of Thoracic and Cardiovascular Surgery. , (2018).</li><li>Townsend, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measuring+Pressure+Volume+Loops+in+the+Mouse.">Measuring Pressure Volume Loops in the Mouse.</a> Journal of Visualized Experiments. (111), e53810 (2016).</li><li>Ratschiller, T., Deutsch, M. -. 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=Heterotopic+Cervical+Heart+Transplantation+in+Mice.">Heterotopic Cervical Heart Transplantation in Mice.</a> Journal of Visualized Experiments. (102), e52907 (2015).</li><li>Fukunaga, N., Bissoondath, V., Rao, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Submandibular+Gland-preserving+Technique+for+Heterotopic+Cervical+Heart+Transplantation+in+Mice.">Submandibular Gland-preserving Technique for Heterotopic Cervical Heart Transplantation in Mice.</a> Transplantation. 1, (2018).</li><li>Gong, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+Heterotopic+Abdominal+Heart+Transplant+Model.">Mouse Heterotopic Abdominal Heart Transplant Model.</a> Rodent Transplant Medicine. , 107-118 (2014).</li><li>Robinson, N., Souslian, L., Gallegos, R. P., Rivard, A. L., Dalmasso, A. P., Bianco, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+Models+for+Cardiac+Research.">Animal Models for Cardiac Research.</a> Handbook of Cardiac Anatomy, Physiology, and Devices. , 469-491 (2015).</li><li>Bianco, R. W., Gallegos, R. P., Rivard, A. L., Voight, J., Dalmasso, A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+Models+for+Cardiac+Research.">Animal Models for Cardiac Research.</a> Handbook of Cardiac Anatomy, Physiology, and Devices. , 393-410 (2009).</li></ol>

The authors have nothing to disclose.

This manuscript describes a large-animal pre-clinical model of orthotopic heart transplantation. Various small animal models of heterotopic heart transplantation have been successfully used to study the effects of novel treatments to improve organ preservation and decrease ischemia-reperfusion injury11,12,13. Additionally, small animal models permit survival models and long-term follow-up to investigate the development of rejection and cardiac allograft vasculopathy11,12,13. However, most of these novel therapies fail in or never make it to clinical trials. In order to facilitate and streamline clinical translation, a reliable and clinically relevant large-animal model is needed.
This protocol was designed to investigate different treatment and organ preservation strategies to prevent or decrease primary graft dysfunction and ischemia-reperfusion injury. As mentioned above, this model has been used since 1994. Authors previously demonstrated the beneficial effects of hypertonic saline infusion in the donor8 or recipient9 prior to organ procurement or implant, respectively. Furthermore, authors investigated different preservation protocols and strategies, such as the use of donor shed blood infusions during cold storage6 and the effect of insulin supplementation into the cardioplegic solution5.
The major limitation of the technique described here is the short-term follow-up. A long-term survival porcine heart transplant model would be resource-intense and involve high-costs. The procedure described here focuses on acute ischemia-reperfusion injury and is a reliable pre-clinical method to investigate novel interventions which have been tested and validated in smaller experimental models, such as the murine model. In addition, this technique can easily by adapted for longer-term follow-up experiments. This would involve adequate heparin reversal, animal decannulation, adequate hemostasis, and chest closure.
The porcine heart is often considered the most anatomically similar to the human heart when using large-animal models. As such, it is an ideal platform to perform cardiac surgical research. However, it is important to note that the tissue is typically described as fragile and friable, especially in the right atrium and the pulmonary artery, being prone to tears14. Additionally, the pig heart is considered sensitive to manipulation and prone to arrhythmias, which is why magnesium sulfate must be routinely administered to each animal at the beginning of the experiment. An important difference between the porcine model and clinical heart transplantation is the left hemiazygous vein in the swine, which drains directly into the coronary sinus. This has to be ligated during the recipient procedure to avoid continuous bleeding. Finally, the porcine model is very sensitive to ischemia, which seems appropriate for acute studies in heart transplantation15.
Recipient management following transplantation can be challenging at times. It is important to revise all anastomoses and ensure no bleeding. A particularly troublesome area is around the posterior pulmonary artery. As mentioned above, the porcine tissues and fragile and can easily tear; if this happens, the surgeon can quickly go back on CPB to correct the issue and attempt weaning once again. Ventricular fibrillation usually occurs during initial reperfusion; if this does not resolve with simple defibrillation, pharmacological interventions, such as 2 g of magnesium sulphate or 1 mg/kg of lidocaine, can be administered and a following defibrillation should be applied. Normal sinus rhythm can be easily achieved in under 3 min.
This procedure requires at least one trained surgeon to be performed; further, 3 to 5 experiments are needed to optimize the protocol within each research group. Additionally, the team should allocate one member to exclusively perform animal anesthesia and recipient management as needed (e.g. inotropic support). Due to the important considerations regarding the porcine model described above, the following steps are critical in this procedure: anesthetic induction and intubation (important to avoid prolong hypoxemic periods), cardiac manipulation during assessment, cannulation for cardiopulmonary bypass, and right atrial and pulmonary artery manipulation and anastomosis. However, as these are routine steps performed in clinical practice, they should be carried out with care and attention to detail. Consistency and repetition will lead to an optimized and reliable model for various uses.

Fifty-years following the first successful report, cardiac transplantation remains the gold-standard treatment for eligible patients with advanced heart failure1. Although ischemic times of up to four hours are tolerated adequately, an ischemic time of greater than six hours is associated with inferior outcomes2. Primary graft dysfunction remains the principal cause of early morbidity and mortality following transplantation2,3. The causes of primary graft dysfunction are multifactorial and include the use of marginal organs, recipient pulmonary vascular disease, hyperacute rejection, and ischemia-reperfusion injury sustained at the time of transplantation3.
Multiple studies have investigated novel methods for donor heart preservation to reduce the incidence of primary graft dysfunction4,5,6,7. It is common practice to assess new techniques and treatments in murine models of ischemia-reperfusion injury or heterotopic heart transplantation. Additionally, small animal models permit survival models and long-term follow-up to investigate the development of rejection and cardiac allograft vasculopathy11,12,13. However, most of these strategies fail initial clinical pilot trials or never reach this stage. It is of paramount importance to evaluate new therapies in a clinically relevant large-animal model for efficient and reliable translation of basic studies&#39;&#160;findings.
The porcine heart is often considered the most anatomically similar to the human heart when using large-animal models. As such, it is an ideal platform to perform cardiac surgical research. However, there are several important factors to consider when using a porcine model. First, the tissue is typically described as fragile and friable, especially in the right atrium and the pulmonary artery, being prone to tears14. Additionally, the pig heart is considered sensitive to manipulation and prone to arrhythmias, which is why one should routinely administer an anti-arrythmetic to each animal at the beginning of the experiment. An important anatomical difference between the porcine model and clinical heart transplantation is the left hemiazygous vein in the swine which drains directly into the coronary sinus. This has to be ligated during the recipient procedure to avoid continuous bleeding. Finally, the porcine model is very sensitive to ischemia, but it is still appropriate for acute studies in heart transplantation15.
This manuscript describes a pre-clinical large-animal (porcine) model of orthotopic heart transplantation that has been firmly established and utilized to investigate novel cardioprotective strategies5,6,8,9.

<table border="0" cellpadding="0" cellspacing="0" style="width:572px;" width="573">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Amiodarone</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td style="width:167px;">Purchased from institutional pharmacy</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Angiocath 20G</td>
			<td style="width:71px;">BD</td>
			<td>381704</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Calcium Chloride 1g/10ml</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>Purchased from institutional pharmacy</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Cardioplegia solution</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>This should be chosen at the investigators discretion.&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Cautery Pencil</td>
			<td style="width:71px;">Covidien</td>
			<td>E2515H</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Central Venous Catheter double-lumen</td>
			<td style="width:71px;">Cook Medical</td>
			<td colspan="2">C-UDLM-501J-LSC</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">CPB pack</td>
			<td style="width:71px;">Medtronic</td>
			<td style="width:119px;"></td>
			<td>Custom-made cardiopulmonary bypass perfusion circuit.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">D5W 5% 250ml</td>
			<td style="width:71px;">Baxter</td>
			<td>JB1064</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">DLP Aortic Root Cannula/stabber</td>
			<td style="width:71px;">Medtronic</td>
			<td>12218</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">DLP single-lumen venous cannula (24F or 28F)</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>This should be chosen at the investigators discretion.&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dobutamine</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>Purchased from institutional pharmacy</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Electrode Polyhesive</td>
			<td style="width:71px;">Covidien</td>
			<td>E7507</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">EOPA arterial cannula (17F or 21F)</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>This should be chosen at the investigators discretion.&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Epinephrine</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>Purchased from institutional pharmacy</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Eppendorf Tubes, 1.5 mL</td>
			<td>Sarstedt&#160;</td>
			<td>72.690.001</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Gloves, nitrile, medium</td>
			<td style="width:71px;">Fischer</td>
			<td>27-058-52</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Heparin 1000 IU/ml</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>Purchased from institutional pharmacy</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Ketalean (Ketamine) inj. 100mg/ml, 50ml/vial</td>
			<td style="width:71px;">Health Canada</td>
			<td style="width:119px;"></td>
			<td>Requires health canada approval</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Lidocaine/Xylocaine 1%</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>Purchased from institutional pharmacy</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Magnesium Sulfate 5g/10ml</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>Purchased from institutional pharmacy</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Midazolam inj. USP 5mg/ml vial/10ml</td>
			<td style="width:71px;">Health Canada</td>
			<td style="width:119px;"></td>
			<td>Requires Health canada approval</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">MPS Quest delivery disposable pack</td>
			<td style="width:71px;">Quest medical</td>
			<td>5001102-AS</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NACL 0.9% 1L</td>
			<td style="width:71px;">Baxter</td>
			<td>JB1324</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Organ Bag</td>
			<td>CardioMed</td>
			<td>2990</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pipette Tips, 1 mL</td>
			<td>Fisherbrand</td>
			<td>02-707-405</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Propofol 1mg/ml</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>Purchased from institutional pharmacy</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Rocuronium</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>Purchased from institutional pharmacy</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Set Admin Prim NF PB W/Checkvalve</td>
			<td style="width:71px;">Smith Medical</td>
			<td style="width:119px;">21-0442-25</td>
			<td>Intravenous infusion pump line. Researchers should choose infusion lines compatible with the infusion pump available at their facilities</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Set Intro Sheath 8.5FRx 10CM</td>
			<td style="width:71px;">Arrow</td>
			<td style="width:119px;">SI-09880</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sofsilk 0 wax coated</td>
			<td style="width:71px;">Covidien</td>
			<td style="width:119px;">S316</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Solumedrol 500mg/5ml</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>Purchased from institutional pharmacy</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Suction tip</td>
			<td style="width:71px;">Covidien</td>
			<td>8888501023</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Suction Tubing 1/4&#34; x 120&#34;</td>
			<td style="width:71px;">Med-Rx</td>
			<td style="width:119px;">70-8120</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Suture 5.0 Prolene BB</td>
			<td style="width:71px;">Ethicon</td>
			<td style="width:119px;">8580H</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Suture Prolene Blum 4-0 SH 36</td>
			<td style="width:71px;">Ethicon</td>
			<td style="width:119px;">8521H</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sutures 2.0 Prolene Blu M SH</td>
			<td style="width:71px;">Ethicon</td>
			<td style="width:119px;">8523H</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sutures BB 4.0 Prolene&#160;</td>
			<td style="width:71px;">Ethicon</td>
			<td style="width:119px;">8881H</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tracheal Tube, 6.5mm</td>
			<td>Mallinckrodt</td>
			<td>86449</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

This pre-clinical model has been used successfully since 19945,6,8,9. Table 1 demonstrates representative results from pressure-volume relationships and echocardiographic parameters taken at baseline, and 3 h post-transplantation in a set of 5 experiments. Although we see a decrease in myocadial contractility following transplantation, this was not statistically significant.
Figure 1 shows representative pressure-volume loops collected from one experiment at the same time-points. During &#34;steady-state&#34;&#160;assessments (Figure 1, top row). volume-dependent parameters are recorded, such as maximum and minimum rate of developed pressure. Volume-independent parameters are obtained by intermittent occlusion of the IVC. With this, the volume of the left ventricle progressively decreases and different relationships can be calculated. In the middle row of Figure 1, we see the end-systolic and end-diastolic pressure-volume relationships being recorded, which represent the relationship between the end-systolic or end-diastolic pressures, respectively, with the corresponding end-diastolic volume. In the bottom row od Figure 1, we see the recording of preload recruitable stroke work, which is the relationship between the stroke work and the corresponding end-diastolic volume.
Finally, as seen in Figure 2, various other metabolic (e.g., lactate levels and pH) and functional parameters (e.g., cardiac output) can be measured with this model to test different hypotheses.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59197/59197fig1.jpg" /> 
Figure 1. Representative pressure-volume loops in a steady state, during Interior Vena Cava (IVC) occlusion, and relationships (preload recruitable stroke work). (A) One experiment at baseline. (B) One experiment following 3 h of reperfusion. PRSW = preload recruitable stroke work. <a href="https://www.jove.com/files/ftp_upload/59197/59197fig1large.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/59197/59197fig2.jpg" /> 
Figure 2. Lactate and pH trends during the heart transplantation protocol. Following reperfusion, there is a significant increase in lactate and decrease in pH. This can be managed by maintaining adequate perfusion pressure with proper volume replacement and vasoactive drug use. <a href="https://www.jove.com/files/ftp_upload/59197/59197fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table>
	<tbody>
		<tr>
			<td></td>
			<td>Baseline</td>
			<td>3 hours post-transplant</td>
			<td>p-value</td>
		</tr>
		<tr>
			<td>Pulmonary artery catheter</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cardiac Index (L/min)</td>
			<td>3.7 &#177; 0.8</td>
			<td>2.8 &#177; 0.3</td>
			<td>0.485</td>
		</tr>
		<tr>
			<td>Pressure-volume analysis</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PRSW (erg&#8729;cm-3&#8729;103) 
			Max dP/dt (mmHg&#8729;s-1) 
			Min dP/dt (mmHg&#8729;s-1)</td>
			<td>62.1 &#177; 7 
			2500 &#177; 425 
			-1537 &#177; 238</td>
			<td>53.8 &#177; 10 
			1815 &#177; 410 
			-1427 &#177; 317</td>
			<td>0.841 
			0.309 
			0.547</td>
		</tr>
		<tr>
			<td>Echocardiography</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>LV EF (%) 
			LV FAC (%) 
			RV FAC (%)</td>
			<td>47.3 &#177; 3.0 
			53.8 &#177; 3.6 
			39.2 &#177; 1.3</td>
			<td>37.0 &#177; 4.2 
			46.4 &#177; 2.9 
			32.8 &#177; 3.6</td>
			<td>0.095 
			0.222 
			0.309</td>
		</tr>
	</tbody>
</table>
Table 1. Representative pressure-volume relationships and echocardiographic parameters from a set of 5 transplants performed at baseline and following 3 h of reperfusion. Data presented as mean &#177; standard error and compared using the Wilcoxon Signed Rank Test. EF = ejection fraction. FAC = fractional area change. LV = left ventricle. Max dP/dt = maximum rate of pressure change in the left ventricle. Min dP/dt = minimum rate pf pressure change in the left ventricule. PRSW = preload recruitable stroke work. RV = right ventricle.

Watch this Scientific Journal Video about A Pre-Clinical Porcine Model of Orthotopic Heart Transplantation at JoVE.com

A Pre-Clinical Porcine Model of Orthotopic Heart Transplantation

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-pre-clinical-porcine-model-of-orthotopic-heart-transplantation

Universidad San Sebastian

Research

JoVE Journal

Medicine

9.4K Views.  University Health Network. 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.

Video: A Pre-Clinical Porcine Model of Orthotopic Heart Transplantation

Orthotopic Aortic Transplantation: A Rat Model to Study the Development of Chronic Vasculopathy

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant vasculopathy (TVP).
The commonly used animal model for cardiovascular chronic rejection studies is the heterotopic heart transplant model performed in laboratory rodents. This model is used widely in experiments since Ono and Lindsey (3) published their technique. To analyze the findings in the blood vessels, the heart has to be sectioned and all vessels have to be measured.
Another method to investigate chronic rejection in cardiovascular questionings is the aortic transplant model (1, 2). In the orthotopic aortic transplant model, the aorta can easily be histologically evaluated (2). The PVG-to-ACI model is especially useful for CAV studies, since acute vascular rejection is not a major confounding factor and Cyclosporin A (CsA) treatment does not prevent the development of CAV, similar to what we find in the clinical setting (4). A7-day period of CsA is required in this model to prevent acute rejection and to achieve long-term survival with the development of TVP.
This model can also be used to investigate acute cellular rejection and media necrosis in xenogeneic models (5).

This video demonstrates the orthotopic aortic transplant model as a simple model to study the development of transplant vasculopathy (TVP) in rats.

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant ...

Development of Obliterative Bronchiolitis in a Murine Model of Orthotopic Lung Transplantation

Orthotopic lung transplantation in rats was first reported by Asimacopoulos and colleagues in 1971 1. Currently, this method is well accepted and standardized not only for the study of allo-rejection but also between syngeneic strains for examining mechanisms of ischemia-reperfusion injury after lung transplantation. Although the application of the rat and other large animal model 2 contributed significantly to the elucidation of these studies, the scope of those investigations is limited by the scarcity of knockout and transgenic rats. Due to no effective therapies for obliterative bronchiolitis, the leading cause of death in lung transplant patients, there has been an intensive search for pre-clinical models that replicate obliterative bronchiolitis. The tracheal allograft model is the most widely used and may reproduce some of the histopathologic features of obliterative bronchiolitis 3. However, the lack of an intact vasculature with no connection to the recipient's conducting airways, and incomplete pathologic features of obliterative bronchiolitis limit the utility of this model 4. Unlike transplantation of other solid organs, vascularized mouse lung transplants have only recently been reported by Okazaki and colleagues for the first time in 2007 5. Applying the basic principles of the rat lung transplant, our lab initiated the obliterative bronchiolitis model using minor histoincompatible antigen murine orthotopic single-left lung transplants which allows the further study of obliterative bronchiolitis immunopathogenesis6.

Obliterative bronchiolitis is the key impediment to the long-term survival of lung transplant recipients and the lack of a robust preclinical model precludes examining obliterative bronchiolitis immunopathogenesis. Unlike other solid organ transplants, vascularized mouse lung transplantation has only recently been developed. Here we show our independently developed obliterative bronchiolitis model after murine orthotopic single-lung transplantation.

Orthotopic lung transplantation in rats was first reported by Asimacopoulos and colleagues in 1971 1. Currently, this method is well accepted and ...

Surgical Procedures for a Rat Model of Partial Orthotopic Liver Transplantation with Hepatic Arterial Reconstruction

Orthotopic liver transplantation (OLT) in rats using a whole or partial graft is an indispensable experimental model for transplantation research, such as studies on graft preservation and ischemia-reperfusion injury 1,2, immunological responses 3,4, hemodynamics 5,6, and small-for-size syndrome 7. The rat OLT is among the most difficult animal models in experimental surgery and demands advanced microsurgical skills that take a long time to learn. Consequently, the use of this model has been limited. Since the reliability and reproducibility of results are key components of the experiments in which such complex animal models are used, it is essential for surgeons who are involved in rat OLT to be trained in well-standardized and sophisticated procedures for this model.
While various techniques and modifications of OLT in rats have been reported 8 since the first model was described by Lee et al. 9 in 1973, the elimination of the hepatic arterial reconstruction 10 and the introduction of the cuff anastomosis technique by Kamada et al. 11 were a major advancement in this model, because they simplified the reconstruction procedures to a great degree. In the model by Kamada et al., the hepatic rearterialization was also eliminated. Since rats could survive without hepatic arterial flow after liver transplantation, there was considerable controversy over the value of hepatic arterialization. However, the physiological superiority of the arterialized model has been increasingly acknowledged, especially in terms of preserving the bile duct system 8,12 and the liver integrity 8,13,14.
In this article, we present detailed surgical procedures for a rat model of OLT with hepatic arterial reconstruction using a 50% partial graft after ex vivo liver resection. The reconstruction procedures for each vessel and the bile duct are performed by the following methods: a 7-0 polypropylene continuous suture for the supra- and infrahepatic vena cava; a cuff technique for the portal vein; and a stent technique for the hepatic artery and the bile duct.

Orthotopic liver transplantation in rats is an indispensable experimental model for biomedical research. Here we present our surgical procedures for orthotopic rat liver transplantation with hepatic arterial reconstruction using a 50% partial graft.

Orthotopic liver transplantation (OLT) in rats using a whole or partial graft is an indispensable experimental model for transplantation research, such as ...

Pre-clinical Evaluation of Tyrosine Kinase Inhibitors for Treatment of Acute Leukemia

Receptor tyrosine kinases have been implicated in the development and progression of many cancers, including both leukemia and solid tumors, and are attractive druggable therapeutic targets. Here we describe an efficient four-step strategy for pre-clinical evaluation of tyrosine kinase inhibitors (TKIs) in the treatment of acute leukemia. Initially, western blot analysis is used to confirm target inhibition in cultured leukemia cells. Functional activity is then evaluated using clonogenic assays in methylcellulose or soft agar cultures. Experimental compounds that demonstrate activity in cell culture assays are evaluated in vivo using NOD-SCID-gamma (NSG) mice transplanted orthotopically with human leukemia cell lines. Initial in vivo pharmacodynamic studies evaluate target inhibition in leukemic blasts isolated from the bone marrow. This approach is used to determine the dose and schedule of administration required for effective target inhibition. Subsequent studies evaluate the efficacy of the TKIs in vivo using luciferase expressing leukemia cells, thereby allowing for non-invasive bioluminescent monitoring of leukemia burden and assessment of therapeutic response using an in vivo bioluminescence imaging system. This strategy has been effective for evaluation of TKIs in vitro and in vivo and can be applied for identification of molecularly-targeted agents with therapeutic potential or for direct comparison and prioritization of multiple compounds.

Receptor tyrosine kinases are ectopically expressed in many cancers and have been identified as therapeutic targets in acute leukemia. This manuscript describes an efficient strategy for pre-clinical evaluation of tyrosine kinase inhibitors for the treatment of acute leukemia.

Receptor tyrosine kinases have been implicated in the development and progression of many cancers, including both leukemia and solid tumors, and are ...

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 ...

Technique of Porcine Liver Procurement and Orthotopic Transplantation using an Active Porto-Caval Shunt

The success of liver transplantation has resulted in a dramatic organ shortage. Each year, a considerable number of patients on the liver transplantation waiting list die without receiving an organ transplant or are delisted due to disease progression. Even after a successful transplantation, rejection and side effects of immunosuppression remain major concerns for graft survival and patient morbidity. 
Experimental animal research has been essential to the success of liver transplantation and still plays a pivotal role in the development of clinical transplantation practice. In particular, the porcine orthotopic liver transplantation model (OLTx) is optimal for clinically oriented research for its close resemblance to human size, anatomy, and physiology. 
Decompression of intestinal congestion during the anhepatic phase of porcine OLTx is important to guarantee reliable animal survival. The use of an active porto-caval-jugular shunt achieves excellent intestinal decompression. The system can be used for short-term as well as long-term survival experiments. The following protocol contains all technical information for a stable and reproducible liver transplantation model in pigs including post-operative animal care.

Experimental animal research plays a pivotal role in the development of clinical transplantation practice. The porcine orthotopic liver transplantation model (OLTx) closely resembles human conditions and is frequently used in clinically oriented research. The following protocol contains all information for a reliable porcine OLTx model using an active porto-caval-jugular shunt.

The success of liver transplantation has resulted in a dramatic organ shortage. Each year, a considerable number of patients on the liver transplantation ...

Pre-clinical Orthotopic Murine Model of Human Prostate Cancer

To study the multifaceted biology of prostate cancer, pre-clinical in vivo models offer a range of options to uncover critical biological information about this disease. The human orthotopic prostate cancer xenograft mouse model provides a useful alternative approach for understanding the specific interactions between genetically and molecularly altered tumor cells, their organ microenvironment, and for evaluation of efficacy of therapeutic regimens. This is a well characterized model designed to study the molecular events of primary tumor development and it recapitulates the early events in the metastatic cascade prior to embolism and entry of tumor cells into the circulation. Thus it allows elucidation of molecular mechanisms underlying the initial phase of metastatic disease. In addition, this model can annotate drug targets of clinical relevance and is a valuable tool to study prostate cancer progression. In this manuscript we describe a detailed procedure to establish a human orthotopic prostate cancer xenograft mouse model.

Prostate cancer is the second most common cause of cancer-related deaths in the United States. An orthotopic cancer model provides a useful approach to understand the biology of prostate cancer and to evaluate the efficacy of therapeutic regimens. This protocol describes detailed steps necessary to establish an orthotopic prostate cancer mouse model.

To study the multifaceted biology of prostate cancer, pre-clinical in vivo models offer a range of options to uncover critical biological information ...

Pre-clinical Model of Cardiac Donation after Circulatory Death

Cardiac transplantation demand is on the rise; nevertheless, organ availability is limited due to a paucity of suitable donors. Organ donation after circulatory death (DCD) is a solution to address this limited availability, but due to a period of prolonged warm ischemia and the risk of tissue injury, its routine use in cardiac transplantation is seldom seen. In this manuscript we provide a detailed protocol closely mimicking current clinical practices in the context of DCD with continuous monitoring of heart function, allowing for the evaluation of novel cardioprotective strategies and interventions to decrease ischemia-reperfusion injury.
In this model, the DCD protocol is initiated in anesthetized Lewis rats by stopping ventilation to induce circulatory death. When systolic blood pressure drops below 30 mmHg, the warm ischemic time is initiated. After a pre-set warm ischemic period, hearts are flushed with a normothermic cardioplegic solution, procured, and mounted onto a Langendorff ex vivo heart perfusion system. Following 10 min of initial reperfusion and stabilization, cardiac reconditioning is continuously evaluated for 60 min using intraventricular pressure monitoring. A heart injury is assessed by measuring cardiac troponin T and the infarct size is quantified by histological staining. The warm ischemic time can be modulated and tailored to develop the desired amount of structural and functional damage. This simple protocol allows for the evaluation of different cardioprotective conditioning strategies introduced at the moment of cardioplegia, initial reperfusion and/or during ex vivo perfusion. Findings obtained from this protocol can be reproduced in large models, facilitating clinical translation.

This protocol shows a simple and flexible approach for the evaluation of new conditioning agents or strategies to increase the feasibility of cardiac donation after circulatory death.

Cardiac transplantation demand is on the rise; nevertheless, organ availability is limited due to a paucity of suitable donors. Organ donation after ...

Orthotopic Kidney Auto-Transplantation in a Porcine Model Using 24 Hours Organ Preservation And Continuous Telemetry

In the present era of organ transplantation with critical organ shortage, various strategies are employed to expand the pool of available allografts for kidney transplantation (KT). Even though, the use of allografts from extended criteria donors (ECD) could partially ease the shortage of organ donors, ECD organs carry a potentially higher risk for inferior outcomes and postoperative complications. Dynamic organ preservation techniques, modulation of ischemia-reperfusion and preservation injury, and allograft therapies are in the spotlight of scientific interest in an effort to improve allograft utilization and patient outcomes in KT.
Preclinical animal experiments are playing an essential role in translational research, especially in the medical device and drug development. The major advantage of the porcine orthotopic auto-transplantation model over ex vivo or small animal studies lies within the surgical-anatomical and physiological similarities to the clinical setting. This allows the investigation of new therapeutic methods and techniques and ensures a facilitated clinical translation of the findings. This protocol provides a comprehensive and problem-oriented description of the porcine orthotopic kidney auto-transplantation model, using a preservation time of 24 hours and telemetry monitoring. The combination of sophisticated surgical techniques with highly standardized and state-of-the-art methods of anesthesia, animal housing, perioperative follow up, and monitoring ensure the reproducibility and success of this model.

Large animal models play an essential role in preclinical transplantation research. Due to its similarities to the clinical setup, the porcine model of orthotopic kidney auto-transplantation described in this article provides an excellent in vivo setting for the testing of organ preservation techniques and therapeutic interventions.

In the present era of organ transplantation with critical organ shortage, various strategies are employed to expand the pool of available allografts for ...

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.