This research is funded on behalf of the University Hospital Foundation.

All the procedures were performed in compliance with the guidelines of the Canadian Council on Animal Care and the guide for the care and use of laboratory animals. The protocols were approved by the institutional animal care committee of the University of Alberta. This protocol has been applied in female juvenile Yorkshire pigs between 35-50 kg. Pigs are pathogen-free, food-grade specimens. They are purchased from the Swine Research and Technology Centre in Edmonton, AB, Canada (https://srtc.ualberta.ca). All individuals involved in ESLP procedures had received proper biosafety training.
1. Pre-surgical preparations and anesthesia
NOTE: Pigs are fasted overnight prior to surgery for a maximum duration of 12 h.
<ol>
	<li>Administer intramuscular injections of ketamine (20 mg/kg) and atropine (0.05 mg/kg) as premedication for the recipient pig in the operating room.</li>
	<li>Place the pig supine on a heated operating table to maintain normothermia and proceed with mask induction.</li>
	<li>Titrate oxygen flow rate according to animal weight and the anesthetic system. 
	NOTE: Oxygen flow should be 20-40 mL/kg.</li>
	<li>Administer isoflurane at 4%-5% and reduce to 3% after 1-2 min.</li>
	<li>Evaluate the depth of anesthesia, ensure the pig has no withdrawal reflex in response to a noxious stimulus. Repeat every 5 min. 
	NOTE: If a pain response is present, increase the percentage of isoflurane administration until the appropriate depth of anesthesia is achieved. See step 10 of this section for further details on maintenance analgesia with ketamine and hydromorphone. No paralytics are administered. This allows for assessment of a withdrawal reflex. A nose pinch is used as a noxious stimulus.</li>
	<li>Intubate the pig once the correct depth of anesthesia is confirmed. Use a custom 10 inch, flat blade laryngoscope and size 9 or 10 endotracheal tubes for pigs 40-50 kg.</li>
	<li>Place a pulse oximeter probe on the tongue (preferred) or ear and target an oxygen saturation above 90%. 
	NOTE: Temperature is monitored via a nasal probe. A heating pad is used to maintain normothermia.</li>
	<li>To maintain the anesthesia, adjust oxygen flow (20-40 mL/kg) and inhalant gas rate (1%-3%).</li>
	<li>Keep the ventilator settings at a respiratory rate of 12-30 breaths/min, TV of 6-10 mL/kg, PEEP of 5 cm H2O, and Peak Pressure of 20 cm H2O. 
	NOTE: A standard ICU style positive pressure ventilator is used to create a closed system for anesthesia and ventilation. Vitals are continuously monitored and recorded at 15 min intervals. ABG&#8217;s are drawn every 15-60 min, depending on the stability of the animal. Although TVs are targeted as high as 10 mL/kg, 6-8 mL/kg are achieved. Figure 1 provides a schematic overview of the negative pressure ventilation (NPV)-ESLP for the transplant protocol applied in the lab.</li>
	<li>Shave, wash and aseptically prepare the incision site using povidone iodine. 
	&#8203;NOTE: Following sedation with Ketamine/Atropine, the analgesic regime involves administering 3mg/kg Ketamine IV q 1 h (range 1-3 mg/kg depending on patient parameters) and Hydromorphone 0.05 mg/kg IM q 2 h via a peripherally inserted IV line in an ear vein. Any longer duration between doses results in breakthrough pain response, such as elevated heart rate and abnormal breathing patterns / abdominal muscle movement.</li>
</ol>
2. Insertion of central venous and arterial lines
<ol>
	<li>Insert a central line for fluid and heparin administration. 
	NOTE: Total IV fluid administration is calculated to 1 mL/kg/h, and fluid boluses are administered PRN to maintain a MAP &#62;60 mmHg. Central line is also used to administer steroids, antibiotics, vasopressors, and inotropes. See Figure 2A for line positioning.
	<ol>
		<li>Prep the skin using a povidone iodine prep solution and allow to dry completely. Use electrocautery to make a 5-8 cm midline incision centered over the trachea and extend cranially from the sternal notch.</li>
		<li>Divide the skin and subcutaneous fat using cautery.</li>
		<li>Divide the midline plane between the strap muscles, and then divide the connective tissue layers to identify the left or right carotid intravascular bundle lateral to the trachea.</li>
		<li>Obtain proximal and distal control of the jugular vein using silk ties (size 2-0) as vessel loops.</li>
		<li>Tie the cranial encircling tie and retract upwards on the proximal tie to control blood flow.</li>
		<li>Make a small incision in the vein using Metzenbaum scissors (see Table of Materials) to accommodate a two-port, 7 Fr central line (~1/3 the vessel&#39;s circumference).</li>
		<li>Simultaneously, release the tension on the proximal vessel loop, cannulate the vein, and then tie down to secure the cannula in the vein at a depth of 10 cm.</li>
		<li>Flush the line with heparin, connect to an IV line of 0.9% normal saline, and administer fluid if the pig is intravascularly depleted from dehydration. 
		NOTE: Heparin lock&#160;any unused ports.</li>
		<li>Administer 500 mg of methylprednisone and 1 g of cefazolin IV.</li>
	</ol>
	</li>
	<li>Follow the same techniques to cannulate the common carotid artery using a 7 Fr arterial line for accurate blood pressure management.</li>
</ol>
3. Left lung procurement
<ol>
	<li>Position the pig in a right lateral decubitus position.</li>
	<li>Perform a left anterolateral thoracotomy (Figure 2).
	<ol>
		<li>Prep the skin using a povidone iodine prep solution and allow to dry completely. Mark the thoracotomy incision (20 cm) using the following landmarks: use palpation to identify the tip of the left scapula; likewise, identify the xiphoid process inferior to the sternum with palpation. Connect the two as shown in Figure 2B.</li>
		<li>Inject a total of 10 mL of 0.25% bupivacaine into the incisional line and two rib spaces above and below the incision.</li>
		<li>Use electrocautery to dissect the skin, subcutaneous layers, and muscle layers. The latissimus dorsi must be divided. Identify the rib immediately below the incision and cauterize on top of the rib to expose the intercostal muscles while avoiding the intercostal neurovascular bundle.</li>
		<li>Use a mosquito hemostat to puncture the intercostal muscles immediately above the rib, and then feel inside the chest for adhesions using a finger. Push the lung away using a Yankauer suction or finger (see Table of Materials) as you cauterize along the top edge of the rib to extend the thoracotomy.
		<ol>
			<li>Extend the thoracotomy anteriorly until 1 inch away from the sternum. Extend the thoracotomy posteriorly to the paraspinal muscles.</li>
		</ol>
		</li>
		<li>Insert a Cooley sternal retractor (see Table of Materials) to open the thoracotomy wide (10 cm) (Figure 2C). Retract the lung to expose the left hemi-azygous vein (Figure 2D).</li>
		<li>Circumferentially dissect the left hemiazygos vein using Metzenbaum scissors and a fine Lauer. Encircle the vessel with silk ties, and then ligate and transect it (Figure 2E). Keep a silk tie on the proximal stump for added control. 
		NOTE: Lauer is a right angle clamp or a celiac clamp used for tissue dissection.</li>
		<li>Dissect out the left pulmonary artery (PA) and left pulmonary veins (PV). Encircle the veins in silk ties for control (Figure 2F). 
		NOTE: The superior PVs are very small and are suture ligated at their branch points or common trunk, depending on the individual anatomy. The left mainstem bronchus is deep to the PA and LA (left atrium), so occasionally, it cannot be dissected easily until the artery and veins have been clamped and transected (Figure 2G).</li>
		<li>Administer 5000 units of heparin IV 5 min before clamping the PA. 
		NOTE: Heparin 5000 units IV is also administered 5 min before unclamping the PA. For every hour after that, 1000 units of IV heparin is administered.</li>
		<li>Clamp the PA (DeBakey cross-clamp), left inferior pulmonary vein (Satinsky clamp), and the left bronchus (Spoon Potts clamp) individually (see Table of Materials). Decrease tidal volumes to 5 mL/kg once the left bronchus is clamped.</li>
		<li>Transect the PA, left inferior pulmonary vein, and the left bronchus. Leave at least 0.5 cm of tissue cuff to sew. Divide the left inferior pulmonary ligament and remove the left lung. 
		NOTE: The left lung can be discarded or kept for control histology.</li>
	</ol>
	</li>
</ol>
4. Termination of ESLP, division of left lung, and flushing with electrolyte solution
<ol>
	<li>Clamp the ventilation tubing at maximal inspiration, terminate perfusion and ventilation, and disconnect the lungs from the ESLP device.</li>
	<li>Weigh the lungs to determine the amount of edema formation. 
	NOTE: Edema is tissue swelling due to the accumulation of excess fluid.</li>
	<li>Take a tissue biopsy of the accessory lobe, divide it into three equal pieces, and place one piece into each of the following: optimum cutting temperature (OCT) gel, formalin, and snap freeze in liquid nitrogen. 
	NOTE: This step is typically followed in the author&#39;s lab. The samples are then stored for future analysis: OCT and snap-frozen samples are kept in a -80 &#176;C freezer, and formalin-stored samples are placed in a properly sealed container and stored in 4 &#176;C refrigerators. Details of the specific ESLP protocol and tissue analysis are published elsewhere16.</li>
	<li>Divide the left donor lung from the right lung. Leave 1 cm of donor PA, 1 cm of donor bronchus, and adequate donor LA cuff (~0.5 cm circumferentially) to sew to the recipient LA (Figure 2H). Leave the left inferior PV and left superior PVs in continuity with the donor LA wall to facilitate later anastomoses.</li>
	<li>Weigh the left lung.</li>
	<li>Cannulate the donor left PA using a drop sucker connected to an IV line and flush 500 mL of extracellular, low potassium, dextran-based electrolyte preservation solution antegrade through the lung vasculature. Secure the cannula in the PA with a silk tie during the flush, and release when the flush is complete. 
	&#8203;NOTE: The steps mentioned pertain to the specific ESLP device utilized for this work and may not be directly applicable to other devices.</li>
</ol>
5. Left lung transplantation
<ol>
	<li>Insert the donor lung into the recipient&#39;s chest, beginning with the lower lobe. Do not force the lung into place. 
	NOTE: The lower ribcage may need to be lifted upwards to accommodate the donor lung by torquing on the sternal retractor. Ideally, the recipient is a few kilograms larger than the donor to facilitate a size match.</li>
	<li>Perform the bronchial anastomosis first using 4-0 prolene on a TF needle (Figure 2I). 
	NOTE: A running, end-to-end anastomosis works well. Trim any excess length from the two anastomotic ends before sewing to avoid kinking caused by redundant tissue.</li>
	<li>Perform the LA anastomosis second with 6-0 prolene on BV-1 needles using a running, end-to-end anastomosis. Again, trim excess tissue to avoid kinking. 
	NOTE: The LA is friable and benefits from the small BV-1 needle. Horizontal bites on the donor may be required to purchase adequate tissue and correct the size mismatched caused by sewing the donor IPV and SPV to the recipient IPV/LA opening.</li>
	<li>Incorporate the donor SPVs into the inferior PV and LA anastomosis to allow left upper lung lobe venous drainage (Figure 2J). 
	NOTE: The branch superior pulmonary veins (SPVs) are less than 0.5 cm in diameter. The common SPV trunk is variable in length and is not routinely present, making direct anastomosis between the donor and recipient SPVs a poor option.</li>
	<li>Complete the PA anastomosis with 6-0 prolene on BV-1 needles using a running, end-to-end anastomosis. Again, trim excess tissue to avoid kinking.</li>
	<li>Remove the bronchial clamp and increase TVs to target 10 mL/kg.</li>
	<li>Confirm heparinization, administer a potassium shift (40 mg of furosemide, 10 units of insulin, 100 mL of 25% dextrose solution), open the PA clamp partially, de-air, and tie the PA suture. Completely release the PA clamp after 10 min.</li>
	<li>Meanwhile, de-air the LA, tie the sutures, and remove the LA clamp.</li>
	<li>Take a reperfusion blood gas from the central line and a reperfusion tissue biopsy from the left middle lobe. 
	NOTE: To take a tissue biopsy, use a size 0-silk tie to encircle a 1 cm portion of the middle lobe apex, tie-down to ensnare the tissue, and then cut the isolated portion with Metzenbaum scissors. Divide the biopsy into three equal portions and manage as previously described.</li>
	<li>Perform a left and right lung bronchoscopy to assess the bronchial anastomosis and to suction secretions. Insert a bronchoscope into the endotracheal tube using an adaptor connection.
	<ol>
		<li>Connect the scope to suction. Advance the bronchoscope into the left bronchus. Inspect the bronchial anastomosis (Figure 2N). Advance the scope down the bronchioles and suction any fluid. Repeat on the right side. 
		NOTE: Do not allow the oxygen saturation to fall below 90%. If saturations fall below this level, remove the scope and allow the pig a few minutes of uninterrupted ventilation to recover.</li>
	</ol>
	</li>
	<li>Insert a 20 Fr malleable chest tube (Figure 2L), close the thoracotomy in three layers (Figure 2M), and prone the pig as soon as the arterial blood gases (ABGs) are stable (Figure 2O).</li>
	<li>Monitor the pig over 4 h in the prone position. Perform an ABG analysis every 30 min. Administer 1000 units of heparin every hour after reperfusion.
	<ol>
		<li>Take a 10 mL blood sample every hour for centrifugation and enzyme-linked immunosorbent assay (ELISA) analysis of inflammatory markers16. 
		&#8203;NOTE: Centrifugation parameters are detailed later.</li>
	</ol>
	</li>
</ol>
6. Isolated Left Lung Assessment
<ol>
	<li>Position the pig supine and re-prep the sternum using povidone iodine prep solution. Perform a midline sternotomy for final isolated left lung assessment (Figure 2P).</li>
	<li>Open the left pleura using Metzenbaum scissors and take a tissue biopsy from the left lower lobe as previously described (NOTE to step 5.9).</li>
	<li>Open the accessory lobe pleura and dissect out the common vein using Metzenbaum scissors. 
	NOTE: This will be clamped later on.</li>
	<li>Take a blood sample from the LA anastomosis using a 21 G needle. Direct the needle toward the left pulmonary veins and away from the common left atrium or accessory lobe trunk.</li>
	<li>Open the right pleura to create space for the right hilar clamps (see Table of Materials). Dissect the right inferior pulmonary ligament up to the hilum. Ensure that a clamp can be placed around the hilum superiorly, inferiorly, and anteriorly. 
	NOTE: This ensures that the hilum is occluded, and all oxygenation is dependent on the left lung. The right lung will not ventilate at this time, which should be evident by a lack of inflation/deflation with ventilator respirations. The right lower lobe can be lifted out of the chest to accomplish this.</li>
	<li>Clamp the accessory lobe vein using a DeBakey aortic cross-clamp (see Table of Materials) to occlude any accessory lobe drainage into the LA (Figure 2Q).</li>
	<li>Clamp the right hilium and take the following serial blood samples from the left PV anastomosis with a 21 G needle directed toward the left lung: 0 min, 1 min, 2 min, 5 min, and 10 min after clamping. 
	NOTE: Five samples are taken to monitor for any trend in partial pressure of oxygen (PaO2) (Figure 2R). The PaO2 should remain relatively stable to represent proper left lung function. Five samples also provide insurance of a quality assessment if there is an issue with clotting of any samples or a problem arises with ABG analysis.</li>
	<li>Transect the anastomoses, and remove the left lung. Transect the IVC to expedite euthanasia under anesthesia via exsanguination. 
	NOTE: Total anesthesia time for the recipient pig is 8 h.</li>
	<li>Weigh the donor lung to assess for edema formation and inspect it for overall appearance. Inspect the PA, bronchus, and LA cuff for signs of clot or other pathology within the donor lung and the recipient mediastinum.</li>
	<li>Run the final gas analyses, centrifuge the perfusate samples, and store the tissue biopsies as previously described (NOTE to step 4.3). 
	NOTE: The centrifugation settings are: 112 x g, 9 acceleration, 9 deceleration, 4 &#176;C, and 15 min duration.</li>
</ol>

<ol>
	<li>Chambers, D. C., 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+international+thoracic+organ+transplant+registry+of+the+international+society+for+heart+and+lung+transplantation:+Thirty-fifth+adult+lung+and+heart-lung+transplant+report-2018;+focus+theme:+Multiorgan+transplantation.">The international thoracic organ transplant registry of the international society for heart and lung transplantation: Thirty-fifth adult lung and heart-lung transplant report-2018; focus theme: Multiorgan transplantation.</a> The Journal of Heart and Lung Transplantation: The Official Publication of the International Society for Heart Transplantation. 37 (10), 1169-1183 (2018).</li><li>Valapour, 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=OPTN/SRTR+2017+annual+data+report:+Lung.">OPTN/SRTR 2017 annual data report: Lung.</a> American Journal of Transplantation: Official Journal of the American Society of Transplantation and the American Society of Transplant Surgeons. 19, 404-484 (2019).</li><li>Klein, A. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organ+donation+and+utilization+in+the+united+states,+1999-2008.">Organ donation and utilization in the united states, 1999-2008.</a> American Journal of Transplantation: Official Journal of the American Society of Transplantation and the American Society of Transplant Surgeons. 10 (4), 973-986 (2010).</li><li>Kotecha, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Continued+successful+evolution+of+extended+criteria+donor+lungs+for+transplantation.">Continued successful evolution of extended criteria donor lungs for transplantation.</a> The Annals of Thoracic Surgery. 104 (5), 1702-1709 (2017).</li><li>Singh, E., 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=Sequence+of+refusals+for+donor+quality,+organ+utilization,+and+survival+after+lung+transplantation.">Sequence of refusals for donor quality, organ utilization, and survival after lung transplantation.</a> The Journal of Heart and Lung Transplantation. 38 (1), 35-42 (2019).</li><li>Cypel, 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=Normothermic+ex+vivo+lung+perfusion+in+clinical+lung+transplantation.">Normothermic ex vivo lung perfusion in clinical lung transplantation.</a> The New England Journal of Medicine. 364 (15), 1431-1440 (2011).</li><li>Wallinder, 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=Early+results+in+transplantation+of+initially+rejected+donor+lungs+after+ex+vivo+lung+perfusion:+A+case-control+study.">Early results in transplantation of initially rejected donor lungs after ex vivo lung perfusion: A case-control study.</a> European Journal of Cardio-Thoracic Surgery: Official Journal of the European Association for Cardio-Thoracic Surgery. 45 (1), 40-44 (2014).</li><li>Sage, E., 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=Lung+transplantation+from+initially+rejected+donors+after+ex+vivo+lung+reconditioning:+The+french+experience.">Lung transplantation from initially rejected donors after ex vivo lung reconditioning: The french experience.</a> European Journal of Cardio-Thoracic Surgery: Official Journal of the European Association for Cardio-Thoracic Surgery. 46 (5), 794-799 (2014).</li><li>Valenza, F., 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=Extracorporeal+lung+perfusion+and+ventilation+to+improve+donor+lung+function+and+increase+the+number+of+organs+available+for+transplantation.">Extracorporeal lung perfusion and ventilation to improve donor lung function and increase the number of organs available for transplantation.</a> Transplantation Proceedings. 44 (7), 1826-1829 (2012).</li><li>Fildes, J. E., 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=Clinical+outcome+of+patients+transplanted+with+marginal+donor+lungs+via+ex+vivo+lung+perfusion+compared+to+standard+lung+transplantation.">Clinical outcome of patients transplanted with marginal donor lungs via ex vivo lung perfusion compared to standard lung transplantation.</a> Transplantation. 99 (5), 1078-1083 (2015).</li><li>Cypel, 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=Experience+with+the+first+50+ex+vivo+lung+perfusions+in+clinical+transplantation.">Experience with the first 50 ex vivo lung perfusions in clinical transplantation.</a> The Journal of Thoracic and Cardiovascular Surgery. 144 (5), 1200-1206 (2012).</li><li>Clark, S. C., 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+new+porcine+model+of+reperfusion+injury+after+lung+transplantation.">A new porcine model of reperfusion injury after lung transplantation.</a> Laboratory Animals. 33, 135-142 (1999).</li><li>Karimi, 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=Technical+pearls+for+swine+lung+transplantation.">Technical pearls for swine lung transplantation.</a> Journal of Surgical Research. 171, 107-111 (2011).</li><li>Kruger, 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=Porcine+pulmonary+auto-transplantation+for+ex+vivo+therapy+as+a+model+for+new+treatment+strategies.">Porcine pulmonary auto-transplantation for ex vivo therapy as a model for new treatment strategies.</a> Interactive CardioVascular and Thoracic Surgery. 23, 358-366 (2016).</li><li>Mariscal, 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=Pig+lung+transplant+survival+model.">Pig lung transplant survival model.</a> Nature Protocols. 13, 1814-1828 (2018).</li><li>Aboelnazar, N. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Negative+pressure+ventilation+decreases+inflammation+and+lung+edema+during+normothermic+ex+vivo+lung+perfusion.">Negative pressure ventilation decreases inflammation and lung edema during normothermic ex vivo lung perfusion.</a> The Journal of Heart and Lung Transplantation: The Official Publication of the International Society for Heart Transplantation. 37 (4), 520-530 (2018).</li></ol>

DHF holds patents on Ex situ organ perfusion technology and methods. DHF and JN are founders and major shareholders of Tevosol, Inc.

Several critical surgical steps are involved in this protocol, and troubleshooting is needed to ensure successful transplantation and lung assessment. Juvenile porcine lungs are incredibly delicate compared to adult human lungs, so the operating surgeon must be cautious when handling porcine lungs. This is especially true after a 12 h run of ESLP as the organ will have taken on fluid volume and be susceptible to injury from excessive manipulation. Any undue pressure will cause atelectasis or trauma to the experimental lu...

Lung transplantation is the preeminent long-term treatment for end-stage lung disease. Over 4,600 lung transplantations are performed worldwide annually1. However, lung transplantation currently has significant limitations. For one, the necessity for organs continues to eclipse available donors. Despite rates of lung transplantation increasing every year since 2012 due to the combined effects of more candidates being listed for transplant, an increase in the number of donors, and improved use of recovered organs, the transplant waitlist mortality has not decreased significantly2. Organ quality concerns represent another major limitation, with reported organ utilization rates as low as 20%-30%3,4,5. Finally, the trends in the post-operative outcomes of lung transplantation are less than satisfactory, with long-term graft and patient outcomes still lagging that of other solid organ transplantations2.
An emerging technology, ex situ lung perfusion (ESLP), has the potential to mitigate these limitations. ESLP has been increasingly applied as a method to assess and recondition marginal donor lungs and has demonstrated acceptable short- and long-term outcomes following transplantation of extended criteria donor (ECD) lungs6,7,8,9,10. Consequently, ESLP has increased utilization rates in some centers by 15%-20%6,7,8,9,10,11.
Proper ESLP research requires the in vivo validation of in vitro findings; however, there is limited literature on porcine lung transplantation models for ESLP12,13,14,15. Furthermore, available literature provides inadequate details regarding anesthetic management of Yorkshire pigs for lung transplantation, which can be highly unstable hemodynamically12,13,14,15. Establishing an easily reproducible model would permit the in vivo validation of current ESLP strategies and the preclinical evaluation of various interventions to reduce lung ischemia-reperfusion injury. The objective of the present study is to describe a porcine model of orthotopic left lung allotransplantation for use with ESLP. The protocol includes descriptions of the anesthetic and surgical techniques, a custom surgical checklist, and details regarding the troubleshooting experience and protocol modifications. The limitations and benefits of the left lung porcine transplantation model have also been discussed in this work. This manuscript does not outline the retrieval process of porcine lungs in 35-50 kg Yorkshire pigs, nor does it cover the establishment and termination of ESLP. This protocol exclusively addresses the recipient transplantation operation.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>ABL 800 FLEX Blood Gas Analyzer</td>
			<td>Radiometer</td>
			<td>989-963</td>
			<td></td>
		</tr>
		<tr>
			<td>Adult-Pediatric Electrostatic Filter HME - Small</td>
			<td>Covidien</td>
			<td>352/5877</td>
			<td></td>
		</tr>
		<tr>
			<td>Allison Lung Retractor</td>
			<td>Pilling</td>
			<td>341679</td>
			<td></td>
		</tr>
		<tr>
			<td>Arterial Filter</td>
			<td>SORIN GROUP</td>
			<td>01706/03</td>
			<td></td>
		</tr>
		<tr>
			<td>Backhaus Towel Clamp</td>
			<td>Pilling</td>
			<td>454300</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>MP biomedicals</td>
			<td>218057791</td>
			<td></td>
		</tr>
		<tr>
			<td>Biomedicus Pump</td>
			<td>Maquet</td>
			<td>BPX-80</td>
			<td></td>
		</tr>
		<tr>
			<td>Bronchoscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cable Ties &#8211; White 12&#8221;</td>
			<td>HUASU International</td>
			<td>HS4830001</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium Chloride</td>
			<td>Fisher Scientific</td>
			<td>C69-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Cooley Sternal Retractor</td>
			<td>Pilling</td>
			<td>341162</td>
			<td></td>
		</tr>
		<tr>
			<td>CUSHING Gutschdressing Forceps</td>
			<td>Pilling</td>
			<td>466200</td>
			<td></td>
		</tr>
		<tr>
			<td>Debakey-Metzenbaum Dissecting</td>
			<td>Pilling</td>
			<td>342202</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>Pilling</td>
			<td>342202</td>
			<td></td>
		</tr>
		<tr>
			<td>DeBakey Peripheral Vascular Clamp</td>
			<td>Pilling</td>
			<td>353535</td>
			<td></td>
		</tr>
		<tr>
			<td>Debakey Straight Vascular Tissue Forceps</td>
			<td>Pilling</td>
			<td>351808</td>
			<td></td>
		</tr>
		<tr>
			<td>D-glucose</td>
			<td>Sigma-Aldrich</td>
			<td>G5767-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Drop sucker</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Endotracheal Tube 9.0mm CUFD</td>
			<td>Mallinckrodt</td>
			<td>9590E</td>
			<td></td>
		</tr>
		<tr>
			<td>Flow Transducer</td>
			<td>BIO-PROBE</td>
			<td>TX 40</td>
			<td></td>
		</tr>
		<tr>
			<td>Infusion Pump</td>
			<td>Baxter</td>
			<td>AS50</td>
			<td></td>
		</tr>
		<tr>
			<td>Inspire 7 M Hollow Fiber Membrane Oxygenator</td>
			<td>SORIN GROUP</td>
			<td>K190690</td>
			<td></td>
		</tr>
		<tr>
			<td>Intercept Tubing Connector 3/8&#34; x 1/2&#34;</td>
			<td>Medtronic</td>
			<td>6013</td>
			<td></td>
		</tr>
		<tr>
			<td>Intercept Tubing 1/4&#34; x 1/16&#34; x 8&#39;</td>
			<td>Medtronic</td>
			<td>3108</td>
			<td></td>
		</tr>
		<tr>
			<td>Intercept Tubing 3/8&#34; x 3/32&#34; x 6&#39;</td>
			<td>Medtronic</td>
			<td>3506</td>
			<td></td>
		</tr>
		<tr>
			<td>Laryngoscope</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Custom-made with 10-inch blade</td>
		</tr>
		<tr>
			<td>Metzenbaum Dissecting Scissors</td>
			<td>Pilling</td>
			<td>460420</td>
			<td></td>
		</tr>
		<tr>
			<td>Medical Carbon Dioxide Tank</td>
			<td>Praxair</td>
			<td>5823115</td>
			<td></td>
		</tr>
		<tr>
			<td>Medical Oxygen Tank</td>
			<td>Praxair</td>
			<td>2014408</td>
			<td></td>
		</tr>
		<tr>
			<td>Medical Nitrogen Tank</td>
			<td>Praxair</td>
			<td>NI M-K</td>
			<td></td>
		</tr>
		<tr>
			<td>Mosquito Clamp</td>
			<td>Pilling</td>
			<td>181816</td>
			<td></td>
		</tr>
		<tr>
			<td>Harken Auricle Clamp</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Organ Chamber</td>
			<td>Tevosol</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PlasmaLyte A</td>
			<td>Baxter</td>
			<td>TB2544</td>
			<td></td>
		</tr>
		<tr>
			<td>Poole Suction Tube</td>
			<td>Pilling</td>
			<td>162212</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Phosphate</td>
			<td>Fischer Scientific</td>
			<td>P285-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>PERFADEX Plus</td>
			<td>XVIVO</td>
			<td>19811</td>
			<td></td>
		</tr>
		<tr>
			<td>Satinsky Clamp</td>
			<td>Pilling</td>
			<td>354002</td>
			<td></td>
		</tr>
		<tr>
			<td>Scale</td>
			<td>TANITA</td>
			<td>KD4063611</td>
			<td></td>
		</tr>
		<tr>
			<td>Silicon Support Membrane</td>
			<td>Tevosol</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Bicarbonate</td>
			<td>Sigma-Aldrich</td>
			<td>792519-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride 0.9%</td>
			<td>Baxter</td>
			<td>JB1324</td>
			<td></td>
		</tr>
		<tr>
			<td>Sorin XTRA Cell Saver</td>
			<td>SORIN GROUP</td>
			<td>75221</td>
			<td></td>
		</tr>
		<tr>
			<td>Sternal Saw</td>
			<td>Stryker</td>
			<td>6207</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Electrocautery Device</td>
			<td>Kls Martin</td>
			<td>ME411</td>
			<td></td>
		</tr>
		<tr>
			<td>TruWave Pressure Transducer</td>
			<td>Edwards</td>
			<td>VSYPX272</td>
			<td></td>
		</tr>
		<tr>
			<td>Two-Lumen Central Venous Catheter 7fr X2</td>
			<td>Arrowg+ard</td>
			<td>CS-12702-E</td>
			<td></td>
		</tr>
		<tr>
			<td>Vorse Tubing Clamp</td>
			<td>Pilling</td>
			<td>351377</td>
			<td></td>
		</tr>
		<tr>
			<td>Willauer-Deaver Retractor</td>
			<td>Pilling</td>
			<td>341720</td>
			<td></td>
		</tr>
		<tr>
			<td>Yankauer Suction Tube</td>
			<td>Pilling</td>
			<td>162300</td>
			<td></td>
		</tr>
		<tr>
			<td>0 ETHIBOND Green 1X36&#34; Endo Loop 0</td>
			<td>ETHICON</td>
			<td>D8573</td>
			<td></td>
		</tr>
		<tr>
			<td>0 PDS II CP-1 2x27&#8221;</td>
			<td>ETHICON</td>
			<td>Z467H</td>
			<td></td>
		</tr>
		<tr>
			<td>1 VICRYL MO-4 1x18&#8221;</td>
			<td>ETHICON</td>
			<td>J702D</td>
			<td></td>
		</tr>
		<tr>
			<td>2-0 SILK Black 12&#34; x 18&#34; Strands</td>
			<td>ETHICON</td>
			<td>SA77G</td>
			<td></td>
		</tr>
		<tr>
			<td>4-0 PROLENE Blue TF 1x24&#8221;</td>
			<td>ETHICON</td>
			<td>8204H</td>
			<td></td>
		</tr>
		<tr>
			<td>6-0 PROLENE Blue BV 2x30&#8221;</td>
			<td>ETHICON</td>
			<td>M8776</td>
			<td></td>
		</tr>
		<tr>
			<td>21-Gauge Needle</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

left lung orthotopic transplantation in a juvenile porcine model for eslp

Lung transplantation is the gold-standard treatment for end-stage lung disease, with over 4,600 lung transplantations performed worldwide annually. However, lung transplantation is limited by a shortage of available donor organs. As such, there is high waitlist mortality. Ex situ lung perfusion (ESLP) has increased donor lung utilization rates in some centers by 15%-20%. ESLP has been applied as a method to assess and recondition marginal donor lungs and has demonstrated acceptable short- and long-term outcomes following transplantation of extended criteria donor (ECD) lungs. Large animal (in vivo) transplantation models are required to validate ongoing in vitro research findings. Anatomic and physiologic differences between humans and pigs pose significant technical and anesthetic challenges. An easily reproducible transplant model would permit the&#160;in vivo&#160;validation of current ESLP strategies and the preclinical evaluation of various interventions designed to improve donor lung function. This protocol describes a porcine model of orthotopic left lung allotransplantation. This includes anesthetic and surgical techniques, a customized surgical checklist, troubleshooting, modifications, and the benefits and limitations of the approach.

All of the results are in the context of 4 h of reperfusion following 12 h of NPV-ESLP16. During lung explant, there are several clinical outcomes to anticipate (Figure 3). Typically, the pig will remain hemodynamically stable following a successful left lung explantation but may require a low dose infusion of phenylephrine (dose range: 2-10 mg/h) due to a vasodilatory response to surgery. Heart rate should target approximately 100-120 bpm, respiratory rate (RR) 8-30 ...

Watch this Scientific Journal Video about Left Lung Orthotopic Transplantation in a Juvenile Porcine Model for ESLP at JoVE.com

Left Lung Orthotopic Transplantation in a Juvenile Porcine Model for ESLP

This protocol describes a juvenile porcine model of orthotopic left lung allotransplantation designed for use with ESLP research. Focus is made on anesthetic and surgical techniques, as well as critical steps and troubleshooting.

Lung transplantation is the gold-standard treatment for end-stage lung disease, with over 4,600 lung transplantations performed worldwide annually. ...

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Research

JoVE Journal

Medicine

1.6K Views.  University of Alberta. This protocol describes a juvenile porcine model of orthotopic left lung allotransplantation designed for use with ESLP research. Focus is made on anesthetic and surgical techniques, as well as critical steps and troubleshooting.

Video: Left Lung Orthotopic Transplantation in a Juvenile Porcine Model for ESLP

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 ...

Orthotopic Small Bowel Transplantation in Rats

Small bowel transplantation has become an accepted clinical option for patients with short gut syndrome and failure of parenteral nutrition (irreversible intestinal failure). In specialized centers improved operative and managing strategies have led to excellent short- and intermediate term patient and graft survival while providing high quality of life 1,3. Unlike in the more common transplantation of other solid organs (i.e. heart, liver) many underlying mechanisms of graft function and immunologic alterations induced by intestinal transplantation are not entirely known6,7. Episodes of acute rejection, sepsis and chronic graft failure are the main obstacles still contributing to less favorable long term outcome and hindering a more widespread employment of the procedure despite a growing number of patients on home parenteral nutrition who would potentially benefit from such a transplant. The small intestine contains a large number of passenger leucocytes commonly referred to as part of the gut associated lymphoid system (GALT) this being part of the reason for the high immunogenity of the intestinal graft. The presence and close proximity of many commensals and pathogens in the gut explains the severity of sepsis episodes once graft mucosal integrity is compromised (for example by rejection). To advance the field of intestinal- and multiorgan transplantation more data generated from reliable and feasible animal models is needed. The model provided herein combines both reliability and feasibility once established in a standardized manner and can provide valuable insight in the underlying complex molecular, cellular and functional mechanisms that are triggered by intestinal transplantation. We have successfully used and refined the described procedure over more than 5 years in our laboratory 8-11. The JoVE video-based format is especially useful to demonstrate the complex procedure and avoid initial pitfalls for groups planning to establish an orthotopic rodent model investigating intestinal transplantation.

Small bowel transplantation has become an accepted treatment option for patients with irreversible intestinal failure. Our experimental model of orthotopic small bowel transplantation in rats serves as a reliable tool to address underlying immunologic and inflammatory processes that complicate intestinal transplantation.

Small  bowel transplantation has become an accepted clinical option for patients with  short gut syndrome and failure of parenteral nutrition ...

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 ...

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 ...

A Murine Model of Myocardial Ischemia-reperfusion Injury through Ligation of the Left Anterior Descending Artery

Acute or chronic myocardial infarction (MI) are cardiovascular events resulting in high morbidity and mortality. Establishing the pathological mechanisms at work during MI and developing effective therapeutic approaches requires methodology to reproducibly simulate the clinical incidence and reflect the pathophysiological changes associated with MI. Here, we describe a surgical method to induce MI in mouse models that can be used for short-term ischemia-reperfusion (I/R) injury as well as permanent ligation. The major advantage of this method is to facilitate location of the left anterior descending artery (LAD) to allow for accurate ligation of this artery to induce ischemia in the left ventricle of the mouse heart. Accurate positioning of the ligature on the LAD increases reproducibility of infarct size and thus produces more reliable results. Greater precision in placement of the ligature will improve the standard surgical approaches to simulate MI in mice, thus reducing the number of experimental animals necessary for statistically relevant studies and improving our understanding of the mechanisms producing cardiac dysfunction following MI. This mouse model of MI is also useful for the preclinical testing of treatments targeting myocardial damage following MI.

We introduce a surgical method to induce experimental ischemia/reperfusion (I/R) injury to simulate myocardial infarction (MI) in mouse models that allows for more clarity in positioning of the ligation on the left anterior descending artery (LAD) to increase the reproducibility of MI experiments in mice.

Acute or chronic myocardial infarction (MI) are cardiovascular events resulting in high morbidity and mortality. Establishing the pathological mechanisms ...

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

The number of acceptable donor lungs available for lung transplantation is severely limited due to poor quality. Ex-Vivo Lung Perfusion (EVLP) has allowed lung transplantation in humans to become more readily available by enabling the ability to assess organs and expand the donor pool. As this technology expands and improves, the ability to potentially evaluate and improve the quality of substandard lungs prior to transplant is a critical need. In order to more rigorously evaluate these approaches, a reproducible animal model needs to be established that would allow for testing of improved techniques and management of the donated lungs as well as to the lung-transplant recipient. In addition, an EVLP animal model of associated pathologies, e.g., ventilation induced lung injury (VILI), would provide a novel method to evaluate treatments for these pathologies. Here, we describe the development of a rat EVLP lung program and refinements to this method that allow for a reproducible model for future expansion. We also describe the application of this EVLP system to model VILI in rat lungs. The goal is to provide the research community with key information and &#8220;pearls of wisdom&#8221;/techniques that arose from trial and error and are critical to establishing an EVLP system that is robust and reproducible.

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

Ex-Vivo Lung Perfusion (EVLP) has allowed lung transplantation in humans to become more readily available by enabling the ability to assess organs and expand the donor pool. Here, we describe the development of a rat EVLP program and refinements that allow for a reproducible model for future expansion.

The number of acceptable donor lungs available for lung transplantation is severely limited due to poor quality. Ex-Vivo Lung Perfusion (EVLP) has allowed ...

Orthotopic Hind Limb Transplantation in the Mouse

In vivo animal model systems, and in particular mouse models, have evolved into powerful and versatile scientific tools indispensable to basic and translational research in the field of transplantation medicine. A vast array of reagents is available exclusively in this setting, including mono- and polyclonal antibodies for both diagnostic and interventional applications. In addition, a vast number of genotyped, inbred, transgenic, and knock out strains allow detailed investigation of the individual contributions of humoral and cellular components to the complex interplay of an immune response and make the mouse the gold standard for immunological research. 
Vascularized Composite Allotransplantation (VCA) delineates a novel field of transplantation using allografts to replace "like with like" in patients suffering traumatic or congenital tissue loss. This surgical methodological protocol shows the use of a non-suture cuff technique for super-microvascular anastomosis in an orthotopic mouse hind limb transplantation model. The model specifically allows for comparison between established paradigms in solid organ transplantation with a novel form of transplants consisting of various different tissue components. Uniquely, this model allows for the transplantation of a viable vascularized bone marrow compartment and niche that have the potential to exert a beneficial effect on the balance of immune acceptance and rejection. This technique provides a tool to investigate alloantigen recognition and allograft rejection and acceptance, as well as enables the pursuit of functional nerve regeneration studies to further advance this novel field of transplantation.

This novel model for orthotopic hind limb transplantation in the mouse, applying a non-suture cuff technique for super-microvascular anastomosis, provides a powerful tool for in&#160;vivo mechanistic immunological research related to vascularized composite allotransplantation (VCA).

In vivo animal model systems, and in particular mouse models, have evolved into powerful and versatile scientific tools indispensable to basic and ...

A Pre-Clinical Porcine Model of Orthotopic Heart Transplantation

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

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

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

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 Rat Orthotopic Renal Transplantation Model for Renal Allograft Rejection

Renal allograft rejection limits the long-term survival of patients after renal transplantation. Rat orthotopic renal transplantation is an essential model to investigate the mechanism of renal allograft rejection in pre-clinical studies and could aid in the development of novel approaches to improve the long-term survival of renal allografts. Donor kidney implantation in rat orthotopic renal transplantation is commonly performed by end-to-side anastomosis to recipients' aorta and inferior vena cava. In this model, the donor's kidney was implanted using end-to-end anastomosis to the recipients' renal artery and renal vein. The donor's ureter was anastomosed to the recipient's bladder in an end-to-side 'tunnel' method. This model contributes to better healing of ureter-bladder anastomosis and increases the recipients' survival by avoiding interference with blood supply and venous reflux of the lower body. This model can be used to investigate the mechanisms of acute and chronic immune and pathologic rejection of renal allografts. Here, the study describes the detailed protocols of this orthotopic renal transplantation between rats.

The rat orthotopic renal transplantation model contributes to investigating the mechanism of renal allograft rejection. The current model increases the recipients' survival without interference with blood supply and venous reflux of the lower body using an end-to-end anastomosis of kidney implantation and an end-to-side "tunnel" method of ureter-bladder anastomosis.

Renal allograft rejection limits the long-term survival of patients after renal transplantation. Rat orthotopic renal transplantation is an essential ...