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>

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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 lung that will affect assessment results. Likewise, the vascular structures are very delicate in the juvenile pig. It is critical to avoid torsion of the PA clamp as this can cause a tear or dissection of the tissue layers. A tear in the PA will necessitate opening the pericardium to access a more proximal portion of the left PA that can be anastomosed to the implanting lung. A DeBakey vascular clamp has a low profile that fits well in the surgical field, but this instrument can cause injury to the delicate PA if the surgeon is not careful. It is helpful to secure the clamp in position using a silk tie that is snapped to the drapes to prevent dislodgement or torsion. Bronchoscopy of the transplanted lung after unclamping of the bronchial anastomosis is also critical. There is often fluid within the donor lung airway after 12 h of ESLP and transplant. Suctioning this fluid is vital to ensure optimal recovery of left lung function and thereby assessment after 4 h of reperfusion. After bronchoscopy and the first ABG has returned with satisfactory potassium levels, it is critical to insert a chest tube, close the incision, and prone the pig. The pig&#39;s hemodynamics and ventilation are considerably more stable in the prone position, with the ribcage reapproximated. Elevated potassium &#62; 5.5 mmol/L at this stage risks bradycardic arrest and will require emergent re-opening and manual cardiac massage to support perfusion, which is best avoided. Due to the significant risk of hyperkalemia and bradycardic arrest upon reperfusion, it is critical to perform serial ABGs beginning at reperfusion and recurring every 30 min until the 4 h exsanguination. ABGs give essential readings of oxygenation, partial pressure of carbon dioxide (PCO2), potassium, and glucose. Monitoring these four components closely and treating them appropriately is vital to a successful experiment. A continuous telemetry reading is also critical to monitor for peaked T waves associated with hyperkalemia and the anticipation of bradycardia. At the final stages of the experiment, it is crucial to clamp the right lung hilum and the accessory lobe before drawing final blood samples from the LA anastomosis. The right hilum supplies blood to the accessory lung lobe, and the accessory lobe drains adjacent to the left inferior pulmonary vein, often via a common trunk. The right hilum and accessory lobe need to be clamped separately to ensure no right lung function contributes to the sample LA gases through blood mixing. Drawing the left lung ABG sample from the PV anastomosis or just beyond it is suggested.
Several modifications have been made to this protocol along with significant troubleshooting of the described methods. Initially, it was attempted to perform the implantation via median sternotomy; however, the exposure was suboptimal due to the orientation of the pig PA, bronchus, and LA. The approach was successfully performed, but a thoracotomy was attempted on subsequent surgeries for improved exposure. This proved to be a superior surgical approach from visualization and technical perspective. Another essential modification was developing and implementing a surgical safety/protocol checklist (Supplementary File 1). There was a significant learning curve for all the team members involved, and these experiments are resource-intensive. A checklist was developed to guide the communication and document protocol development (Supplementary File 1). The checklist allowed to systemize and simplify the protocol for faster learning. The heparinization protocol was also modified. Two of the first ten transplants performed suffered from left lung ischemia due to clot formation in the left PA. Initially, 5000 units of heparin IV was administered 5 min before PA clamping and an additional 5000 units was administered 5 min before PA unclamping. Dosing frequency was increased to include 5000 units every hour after PA unclamping, and there have not been any issues with bleeding or PA clotting since adopting this approach. A strategy that utilizes less heparin was developed to control expenses, with a dose of 5000 units IV heparin 5 min before PA clamping and 5 min before partial PA unclamping. This is followed by 1000 unit IV heparin boluses every hour for the remainder of the case. There was no access to ACT analysis, which would be the most accurate means of accessing adequacy of heparinization.
The unclamping of the PA was also modified from a sudden unclamping to an approach that gradually reintroduces full flow to the transplanted lung over 10 min. The left inferior PV and LA cuff remain clamped upon PA unclamping to allow for antegrade de-airing. Full PA flow produced significant pressure on the delicate LA suture lines and considerable pressure within the lung vasculature, which appeared damaging. Prolonged PA unclamping allows for the antegrade de-airing of the LA with a gradual increase in flow as opposed to sudden unclamping and a sudden increase in flow. Prolonged unclamping protects the suture lines and lung endothelium from sudden increase in pressure. Even with ESLP, an ischemic insult to the transplanted lung and cell death contributes to a significant release of potassium into the pig&#39;s circulation following ischemic-reperfusion. For managing hyperkalemia proactively, the protocol was modified to pre-emptively shift potassium at the time of reperfusion by administering furosemide 40 mg IV, 100 mL of 25% dextrose (D25), and 10 units of regular insulin. This maintains target potassium on the ABGs within the first hour of reperfusion, and the pig can be safely proned earlier in the experiment, which helps with graft function. From a hemodynamic perspective, the protocol is modified to use phenylephrine as the predominant vasopressor support. Vasopressin was found to be less effective. A low dose drip of dobutamine was occasionally run to increase cardiac output, along with a phenylephrine infusion to maintain blood pressure. Still, dobutamine is used sparingly due to its arrhythmogenic properties. Finally, the assessment of the isolated left lung was modified. After clamping the right lung hilum, the LA gases were initially drawn from the body of the LA after lifting the heart cephalad; however, gas mixing from the accessory lobe drainage into the LA produced falsely high PaO2 readings. Now, samples are drawn distal to the LA anastomosis line after clamping the right lung and the accessory lobe individually. These samples are taken at 0, 1, 2, 5, and 10 min after clamping the right hilum and are a more accurate representation of the isolated left lung function. Manual cardiac massage may be required between the 5-10 min mark. The most recent protocol improvement pertains to the superior pulmonary vein (SPV) anastomoses. Initially, the recipient SPVs were oversewn due to their small caliber and propensity to clot.&#160;Still, the donor&#39;s upper lobe occasionally suffered congestion as collateral drainage was variable and inadequate between pigs. To remedy this, the donor SPV and IPV were incorporated into the recipient&#39;s IPV/LA anastomosis, eliminating any issue with venous drainage and lung congestion. This protocol will continue to benefit from further modification as experience grows.
There are several limitations with this method of left lung transplantation.&#160;The model has only been assessed with a 4 h period, which only considers the transplanted lung function in the acute post-operative period following 12 h of ESLP. This protocol was designed with the animal&#39;s recovery in mind; however, it has yet to be tested in that capacity. The technical operation requires considerable surgical skill and necessitates a trained surgeon or highly independent surgical trainee to perform. There are many opportunities for fatal errors to occur that would compromise the entire experiment, and proper surgical technique is needed to avoid or correct such hazards. The only true assessment of the transplanted lung occurs at the very end of reperfusion. The native right lung is capable of meeting the oxygen requirements of the pig and producing satisfactory ABGs. When the right lung is completely clamped at the hilum, it is prevented from receiving fresh oxygen, fresh deoxygenated blood supply, and oxygenated blood drainage. This is a pivotal moment to determine the transplanted left lung function as 100% of cardiac output is redirected toward the transplanted lung, which becomes solely responsible for systemic oxygenation.
There are multiple benefits of this method concerning existing/alternative methods. After reviewing the literature12,13,14,15, this method is the most detailed and reproducible after an initial learning curve of 1 or 2 pigs in the hands of a junior cardiac surgical trainee or fully qualified surgeon. The operation is straightforward; however, the hemodynamics of the pig (including its susceptibility for lethal arrhythmias) creates a learning opportunity for those accustomed to operating on adult humans, which are more robust from a cardiopulmonary perspective. The methods for isolated left lung functional assessment, although brief, are easy to perform and highly reproducible. In particular, this methodology provides more details about anesthetic management than is currently available in the literature.
In vivo transplantation is essential for ESLP and lung transplantation research.&#160;ESLP is the most crucial development in lung transplantation since the introduction of antirejection medication, with some centers already benefitting from the increased organ utilization rates afforded by this technology6,7,8,9,10,11,12. Further advancement in this field of research is needed to decrease waitlist mortality and expand the accessibility of ESLP platforms. In vitro analysis with ESLP benefits from the in vivo assessment and confirmation of a large animal model. Large animal models that confirm in vitro findings are often necessary for clinical research trial approval for developing labs. This method provides a reliable and relatively straightforward transplant method for labs performing ESLP research.

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>
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		<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>
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		<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>
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		<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>
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		<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>
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		<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 for SpO2 &#62; 90%, mean arterial pressure (MAP) &#62; 60 mmHg, normothermic (38 &#176;C), and tidal volumes (TVs) are targeted at 5 mL/kg while on one-lung ventilation with peak pressures of 20-24 cm H2O. During one-lung ventilation, the ventilation volumes were reduced by half to protect the left lung from overinflation. The respiratory rate was increased to target a physiologic end-tidal carbon dioxide level (Figure 3). Thus, Figure 3 displays typical hemodynamic and ventilatory parameters during critical points of the transplant.
During lung implant, the following results are typical. The left lung will have absorbed fluid during the ESLP run and appears heavier and larger than the explanted lung. For this reason, the recipient should be slightly larger than the donor (2-4 kg), so the thorax can accommodate the somewhat edematous lung. The lung will require gentle pressure to insert into the chest through the thoracotomy. It is easier to insert the lower lobe first, followed by the upper lobe. The bronchus is a direct end-to-end anastomosis and should be performed first. 4-0 prolene on a TF needle is recommended. The LA cuffs are highly friable but not too difficult to sew due to the redundancy and pliability of the tissue. 6-0 prolene on BV-1 needles work well for the LA anastomoses. The PA is the last anastomosis performed. This vessel can tear easily with little traction. If it tears, it is possible to open the pericardium and move the clamp proximally toward healthy tissue for sewing. Again, a 6-0 prolene on BV-1 needles works well for this anastomosis.
At the time of reperfusion, the following trends were noticed. Once the bronchus is unclamped and TVs are increased back to 10 mL/kg, the left lung will begin to inflate. Although the target was 10 mL/kg for tidal volumes, generally 6-8 mL/kg was attained, which is achieved gradually over the first 2-3 h of reperfusion, depending on the ESLP protocol used and the quality of the implanted lung. Rarely, there can be a small air leak, and this can be remedied with a simple stitch on the anterior wall. The posterior wall is more difficult to repair and will require packing. Great effort should be made to avoid air leaks from the bronchial anastomosis. Upon bronchoscopy, the right lung appears normal, and the left lung is typically edematous. The suture line is inspected, and approximately 50-100 mL of clear fluid is suctioned from the airways. The TV will drop significantly during suctioning from 300 s to 20 s, so this action should be performed quickly to allow the pig to recover. If arterial saturation drops below 90%, the bronchoscopy should be terminated, and the pig is allowed to recover over 1-2 min of ventilation. The first arterial blood gas (ABG) is typically normal because the right lung is functioning well as the left lung recovers.
The proactive administration of furosemide, dextrose, and insulin at the time of reperfusion serves to mitigate a dramatic rise in potassium through intracellular shifting. The potassium will predictably rise during 60-120 min of reperfusion (Table 1). Table 1 demonstrates a sample of ABGs over transplantation with 4 h reperfusion following 12 h of normothermic negative pressure ventilation (NPV) ESLP. Approximately two to four shifts are required during 4 h reperfusion to keep potassium &#60; 5 mmol/L. If the trend is upward and appears as a rapid change between two gases drawn at 30 min intervals, the target is K+&#60; 4.5 mmol/L. Shifts include 40 mg of furosemide, 100 mL of 25% dextrose (D25), and 10 units of regular insulin administered as IV push via the central line. Occasionally, the pig will require a low dose dobutamine infusion (1.5-5 mcg/kg/min) along with phenylephrine (2-10 mg/h) after 30-60 min of reperfusion to treat a developing vasoplegic response. It is preferable to use phenylephrine in this situation exclusively. However, dobutamine can be a useful supplemental inotrope to maintain a mean arterial pressure greater than 60 mmHg, particularly if the heart rate is bradycardic.
Upon thoracotomy closure and turning the pig prone, an improvement in ventilation and hemodynamics is demonstrated. The modification can be drastic and occur over 5-10 min, but occasionally the response takes 1 h. Tidal volumes increase as pressure/weight is taken off the right lung, and the left lung continues to ventilate with improved compliance and recruitment. A repeat bronchoscopy can be performed further to clear the airway after a change in position. Over the following 4 h, phenylephrine requirements decrease, TVs approach the target 10 mL/kg, and ABGs stabilize (Table 1). To reiterate, if TVs of 10 mL/kg are targetted, typically TVs in the range of 6-8 mL/kg are achieved (Figure 3).
At the time of the final isolated left lung assessment, a stable pattern of behavior has been observed. The pig is less tolerant hemodynamically in the supine position for sternotomy and may require additional vasopressor support. Inspection of the left lung reveals variable degrees of mild hyperemia from ischemic reperfusion injury (IRI). The right lung appears normal. Upon clamping the right hilum, the pig becomes sinus tachycardic (120-140 bpm), and 100% of the cardiac output is diverted to the left lung. Targeted tidal volumes are not decreased at this time as the entire process takes 10 min. The pig remains stable up to the 5 min mark, but the heart may develop ventricular fibrillation between 5-10 min and manual cardiac massage&#160;is&#160;potentially required to continue perfusing the left lung. The left lung is explanted, weighed, and the anastomoses are inspected for patency. The pig expires rapidly at the time of exsanguination, which coincides with the explantation of the previously transplanted lung.
A successful transplant has predictable findings after the experiment (Table 1 and Figure 4). Figure 4 displays typical P:F ratio changes and edema formation during the transplant protocol. Typically, the left lung will experience an approximate 35% (+/-15%) weight gain; however, residual blood in the circulation contributes to this weight. PF ratios drop by approximately 100 at reperfusion as the left lung is not immediately effective at oxygenation, but this discrepancy improves over 2-3 h. Upon isolated left lung assessment at 4 h, the PF ratio will remain stable or decline slightly. Generally, the isolated left lung gas at 10 min will be similar to the final gas analysis post 12 h ESLP (Table 1). However, this is entirely dependent on the ESLP protocol employed, and the extent of IRI incurred. An unsuccessful transplant can be caused by clotting of the LPA, which results in an infarcted lung that does not oxygenate. Likewise, the duration of the transplant surgery can affect the quality of the reperfused lung function. An implantation surgery should take between 30-60 min. Longer operations expose the donor lung to damaging warm ischemic time that exacerbates ischemic reperfusion injury and can confound the results of the experimental ESLP protocol. The specific ESLP protocol of a given experiment may produce a non-functioning lung that fails to oxygenate after transplantation despite patent anastomoses. Such isolated left lung gases will be very dark in color (deoxygenated) with a low partial pressure of oxygen (PaO2).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62979/62979fig01.jpg" /> 
Figure 1: Schematic of porcine left lung transplant protocol. Schematic representation of 12 h NPV-ESLP run followed by left lung transplantation in a Yorkshire pig. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62979/62979fig01large.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/62979/62979fig02.jpg" /> 
Figure 2: Photos of porcine left lung transplant surgery protocol. (A) Internal jugular and common carotid line placement. (B) Thoracotomy incision. (C) Thoracotomy. (D) Left Hemi-azygous vein. (E) Ligated Left Hemi-azygous vein. (F) Isolation of pulmonary veins. (G) Clamped left atrial cuff, left bronchus, and left pulmonary artery. (H) Left donor lung with pulmonary vein, bronchial and PA cuffs. (I) Pulmonary artery anastomosis. (J) Left lung transplanted and unclamped. (K) Lung repositioned. (L) Chest tube positioned. (M) Thoracotomy closure. (N) Bronchial anastomosis. (O) Pig in prone position. (P) Sternotomy. (Q) Accessory lobe clamped (Right lung clamped, but not shown). (R) Left pulmonary vein blood samples were drawn from pulmonary vein anastomosis (bleeding from prior puncture site). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62979/62979fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62979/62979fig03.jpg" /> 
Figure 3: Monitoring and ventilation parameters for porcine left lung transplant surgery. (A) Typical parameters for recipient pre-transplant. (B) Typical parameters at recipient left lung explant. (C) Typical parameters 4 h post left lung donor transplant. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62979/62979fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62979/62979fig04.jpg" /> 
Figure 4: P:F ratio and weight gain pre-and post-transplant. (A) PaO2:FiO2 ratios throughout the transplant. (B) Weight gain of left lung throughout transplant after 12 h of NPV-ESLP. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62979/62979fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:font-size="7px" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Arterial Blood Gases (100% FiO2)</td>
			<td>In vivo Recipient</td>
			<td>T0 Reperfusion</td>
			<td>T1 Reperfusion</td>
			<td>T2 Reperfusion</td>
			<td>T3 Reperfusion</td>
			<td>T4 Reperfusion</td>
			<td>Isolated Left Lung Pre-clamp</td>
			<td>Isolated Left Lung Post-clamp (0 min)</td>
			<td>Isolated Left Lung Post-clamp (1 min)</td>
			<td>Isolated Left Lung Post-clamp (5 min)</td>
			<td>Isolated Left Lung Post-clamp (10 min)</td>
		</tr>
		<tr>
			<td>Blood Gas Values</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>pH</td>
			<td>7.402</td>
			<td>7.327</td>
			<td>7.284</td>
			<td>7.402</td>
			<td>7.421</td>
			<td>7.479</td>
			<td>7.504</td>
			<td>7.399</td>
			<td>7.371</td>
			<td>7.423</td>
			<td>7.435</td>
		</tr>
		<tr>
			<td>pCO2 (mmHg)</td>
			<td>47.7</td>
			<td>57.3</td>
			<td>56.4</td>
			<td>36.9</td>
			<td>35.3</td>
			<td>35.6</td>
			<td>34.2</td>
			<td>45.6</td>
			<td>48.1</td>
			<td>40.6</td>
			<td>36.6</td>
		</tr>
		<tr>
			<td>pO2 (mmHg)</td>
			<td>299</td>
			<td>184</td>
			<td>165</td>
			<td>355</td>
			<td>358</td>
			<td>300</td>
			<td>327</td>
			<td>287</td>
			<td>207</td>
			<td>335</td>
			<td>249</td>
		</tr>
		<tr>
			<td>Oximetry Values</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hb (g/dL)</td>
			<td>11.2</td>
			<td>12.5</td>
			<td>11.3</td>
			<td>11.6</td>
			<td>10.3</td>
			<td>-</td>
			<td>17.1</td>
			<td>11.7</td>
			<td>13.5</td>
			<td>16.3</td>
			<td>13.8</td>
		</tr>
		<tr>
			<td>sO2 (%)</td>
			<td>100.1</td>
			<td>99.2</td>
			<td>99</td>
			<td>99.8</td>
			<td>99.8</td>
			<td>-</td>
			<td>99.9</td>
			<td>100.2</td>
			<td>99.7</td>
			<td>99.8</td>
			<td>99.9</td>
		</tr>
		<tr>
			<td>Electrolyte Values</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>K+ (mmol/L)</td>
			<td>4.5</td>
			<td>6.2</td>
			<td>4.4</td>
			<td>4</td>
			<td>4.1</td>
			<td>4.6</td>
			<td>5.2</td>
			<td>5.4</td>
			<td>5.3</td>
			<td>6.9</td>
			<td>7.4</td>
		</tr>
		<tr>
			<td>Na+ (mmol/L)</td>
			<td>141</td>
			<td>143</td>
			<td>140</td>
			<td>245</td>
			<td>145</td>
			<td>144</td>
			<td>140</td>
			<td>141</td>
			<td>139</td>
			<td>137</td>
			<td>136</td>
		</tr>
		<tr>
			<td>Ca2+ (mmol/L)</td>
			<td>0.99</td>
			<td>0.88</td>
			<td>0.81</td>
			<td>0.74</td>
			<td>0.66</td>
			<td>0.61</td>
			<td>0.36</td>
			<td>0.98</td>
			<td>0.42</td>
			<td>0.36</td>
			<td>0.38</td>
		</tr>
		<tr>
			<td>Cl- (mmol/L)</td>
			<td>97</td>
			<td>97</td>
			<td>95</td>
			<td>101</td>
			<td>100</td>
			<td>96</td>
			<td>91</td>
			<td>102</td>
			<td>94</td>
			<td>91</td>
			<td>94</td>
		</tr>
		<tr>
			<td>Osm (mmol/kg)</td>
			<td>287</td>
			<td>287.9</td>
			<td>293.7</td>
			<td>292.4</td>
			<td>297.5</td>
			<td>293.5</td>
			<td>284.7</td>
			<td>287.1</td>
			<td>282.9</td>
			<td>278.2</td>
			<td>277.1</td>
		</tr>
		<tr>
			<td>Metabolite values</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose (mmol/L)</td>
			<td>4,2</td>
			<td>2.7</td>
			<td>13.4</td>
			<td>2.8</td>
			<td>8.3</td>
			<td>5</td>
			<td>5.1</td>
			<td>4.9</td>
			<td>4.5</td>
			<td>4.6</td>
			<td>4.2</td>
		</tr>
		<tr>
			<td>Lactate (mmol/L)</td>
			<td>1.2</td>
			<td>1.3</td>
			<td>3.8</td>
			<td>2.5</td>
			<td>1.3</td>
			<td>1.2</td>
			<td>1.4</td>
			<td>1.8</td>
			<td>1.4</td>
			<td>1.9</td>
			<td>2.7</td>
		</tr>
		<tr>
			<td>Acid Base status</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>HCO-3 (mmol/L)</td>
			<td>29</td>
			<td>29.1</td>
			<td>25.9</td>
			<td>22.4</td>
			<td>22.5</td>
			<td>26.1</td>
			<td>26.7</td>
			<td>27.6</td>
			<td>27.1</td>
			<td>26.1</td>
			<td>24.1</td>
		</tr>
	</tbody>
</table>
Table 1: Blood gas analysis performed following left lung transplant post 12 h of ESLP. Ca+, calcium ion; Cl-, chloride ion; Hb, hemoglobin; HCO3-, bicarbonate ion; K+, potassium ion; Na+, sodium ion; Osm, osmolarity; paCO2, arterial partial pressure of carbon dioxide; PaO2, arterial partial pressure of oxygen; sO2, oxygen saturation; isolated left lung pre-clamp, right hilum open; Isolated left lung post-clamp, 1 min after right hilum clamped.
Supplementary File 1: Surgical safety checklist for left lung transplantation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62979/Supplementary File 1.eps" target="_blank">Please click here to download this File.</a>

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 ...

Left Lung Orthotopic Transplantation in a Juvenile Porcine Model for ESLP

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

Orthotopic Small Bowel Transplantation in Rats

Orthotopic Liver Transplantation in Rats

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

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

Orthotopic Hind Limb Transplantation in the Mouse

A Pre-Clinical Porcine Model of Orthotopic Heart Transplantation

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

A Rat Orthotopic Renal Transplantation Model for Renal Allograft Rejection