The authors thank Britta Trautewig, Corinna L&#246;bbert, Astrid Dinkel, and Ingrid Meder for their diligence and commitment. Furthermore, the authors thank Tom Figiel for producing the picture material.

This study was performed at the Laboratory for Animal Science of Hannover Medical School after approval by the Lower Saxony regional authority for consumer protection and food safety (Nieders&#228;chsisches Landesamt f&#252;r Verbraucherschutz und Lebensmittelsicherheit [LAVES]; 19/3146)
1. Donor liver procurement
NOTE: The liver donors were female domestic pigs (Sus scrofa domesticus), aged 4-5 months old and with an average body weight of approximately 50 kg, which had already been in quarantine at the animal research facility for a minimum of 10 days prior to surgery.
<ol>
	<li>Perform premedication by intramuscular injection of atropine (0.04-0.08 mg/kg body weight), zolazepam (5 mg/kg body weight), and tiletamine (5 mg/kg body weight). After establishing an intravenous access (e.g., ear vein) induce anesthesia with an injection of propofol (1.5 - 2.5 mg/kg body weight).</li>
	<li>Perform intubation with an 8.0-8.5 mm endotracheal tube, depending on the animal size and anatomy. Establish monitoring of electrocardiography, measurement of respiratory gases and peripheral oxygen saturation, and non-invasive blood pressure measurement.</li>
	<li>Maintain anesthesia in pigs during donor liver procurement via inhalation of isoflurane (0.8-1.5 vol%) and intravenous application of fentanyl (0.003-0.007 mg/kg body weight). Perform volume-controlled ventilation throughout the procedure.</li>
	<li>After placement of the donor pig in a supine position and fixation of the limbs at the base of the operation table with elastic bands, scrub the skin with antiseptic agent, e.g., povidone-iodine or isopropyl alcohol, and cover the animal with sterile drapes.</li>
	<li>Confirm an adequate depth of anesthesia by loss of the withdrawal response to toe pinch. Perform a midline laparotomy beginning at the xiphoid process by using monopolar cautery. Place an abdominal retractor and mobilize the intestine to the right of the donor.</li>
	<li>Perform a splenectomy by dissection of the splenocolic ligament, the gastrosplenic ligament, and the phrenicosplenic ligament. Clamp the splenic vein and splenic artery near the splenic hilum with an Overholt clamp and place ligatures (3-0 polyfilament suture) after severing the vessels. Sever additional (smaller) vessels either by bipolar forceps or by ligation. 
	NOTE: A splenectomy during donor liver procurement is not obligatory but reduces the efflux of blood during and after perfusion.</li>
	<li>Mobilize the intestine to the left side of the donor and sever the falciform ligament and the triangular ligaments using scissors and bipolar cautery.</li>
	<li>After sufficient dissection of the liver, incise the left portion of the diaphragm over a distance of 5-10 cm using scissors to locate the thoracic segment of the descending aorta. Encircle and place a ligature (3-0 polyfilament suture) without tightening.</li>
	<li>Incise the right portion of the diaphragm over a distance of 5-10 cm using scissors and identify the suprahepatic vena cava inferior.</li>
	<li>Relocate the intestine to the upper left of the donor and enter the retroperitoneal space by transverse incision of the peritoneum over a distance of 5-10 cm using scissors.</li>
	<li>Locate the abdominal aorta and inferior vena cava just above the iliac bifurcation and separate both vessels over a length of approximately 6 cm. Place two 3-0 polyfilament ligatures&#160;around the abdominal aorta: one cranial of the iliac bifurcation and one approximately 3 cm cranially, without tightening. Place another&#160;ligature around the intrahepatic vena cava inferior without tightening.</li>
	<li>Intravenously inject heparin (25,000 I.E.). Choose an appropriate cannula and de-air the drip line with cooled preservation solution.</li>
	<li>Tighten the caudally located first ligature around the abdominal aorta. After occluding the abdominal aorta cranially of the second ligature (either manually or by placing an atraumatic vascular clamp), make a transverse incision between both the ligatures using scissors.</li>
	<li>Insert the cannula into the incision and secure it with the remaining ligature. Sever the suprahepatic inferior vena cava far cranially (close to the right atrium) using scissors.</li>
	<li>After blood loss of approximately 1,500-2,000 mL, cross-clamp the thoracic segment of the descending aorta by tying the ligature and start antegrade perfusion. 
	NOTE: For the possible need for blood (transfusions) during engraftment or for normothermic machine perfusion, whole blood (approximately 1,500 mL) can be collected using a container containing citrate-based anticoagulant.</li>
	<li>Tighten the ligature placed around the infrahepatic vena cava inferior, incise the vessel cranially of the ligature, and insert a surgical aspirator. Inject a lethal dose of pentobarbital sodium (5,000 mg). Place crushed sterile ice into the thoracic and abdominal cavity without compromising the liver tissue.</li>
	<li>After perfusion with 3,500 mL of preservation solution over a course of approximately 10-15 min, sever the incised suprahepatic vena cava inferior. Sever the infrahepatic vena cava inferior at the level of the left renal vein.</li>
	<li>Sever the bile duct cranial of the pancreatic tissue between two ligatures (3-0 polyfilament) to avoid bile spillage. Sever the portal vein cranial of the pancreas.</li>
	<li>Locate the celiac artery after blunt preparation and follow dorsally to the abdominal aorta. Excise the respective aortic segment in order to create a patch for later engraftment.</li>
	<li>Excise the diaphragm around the suprahepatic vena cava inferior and sever remaining adhesions using scissors. Extract the liver.</li>
	<li>Perform a cholecystectomy or tighten a ligature around the cystic duct and flush the common bile duct with at least 20 mL of preservation solution. Place the perfusion cannula into the portal vein and flush the graft with a further 500 mL of preservation solution. Place the graft in a sterile bowl placed on ice. 
	&#8203;NOTE: Depending on the scientific objective, the organ can be immediately prepared for engraftment or kept on ice for an indefinite amount of time (20 h in this protocol) before beginning back-table preparation and engraftment.</li>
</ol>
2. Back-table preparation of the liver
<ol>
	<li>Remove the lymphatic tissue beginning at the aortic segment and thereby identify and occlude the arterial side branches and lymphatic vessels with either clips, ligatures (4-0 polyfilament), or sutures (5-0 monofilament; Figure 1A). Likewise, remove the lymphatic tissue around the portal vein and occlude the side branches with sutures (5-0 monofilament).</li>
	<li>Identify the suprahepatic vena cava inferior and place sutures around both diaphragmatic veins (5-0 monofilament) after removing surrounding diaphragmatic tissue. Flush all the vessels with cold saline or preservation solution to identify any remaining leakages. Perform shortening of the vessels and preparation of the aortic patch only upon engraftment to take into account the individual anatomic circumstances.</li>
</ol>
3. Recipient hepatectomy, donor liver engraftment, and perioperative management
NOTE: As liver recipients, female domestic pigs (Sus scrofa domesticus) aged 4-5 months old and with an average body weight of approximately 50 kg, were used. Analogously to the liver donors, the recipients had been in quarantine at the animal research facility for a minimum of 10 days prior to transplant.
<ol>
	<li>Anesthesia and perioperative management
	<ol>
		<li>Perform premedication by intramuscular injection of atropine (0.04-0.08 mg/kg body weight), zolazepam (5 mg/kg body weight), and tiletamine (5 mg/kg body weight). After establishing an intravenous access (e.g., ear vein), induce anesthesia with an injection of propofol (1.5-2.5 mg/kg body weight).</li>
		<li>Perform intubation with an 8.0-8.5 mm endotracheal tube, depending on the animal size and anatomy. Establish monitoring of electrocardiography, measurement of respiratory gases and peripheral oxygen saturation, and non-invasive blood pressure measurement. In the case of a chronic model, apply eye ointment to avoid dryness after the surgical intervention.</li>
		<li>Place the recipient animal on a heating base in a supine position and fix the limbs on the base of the operation table with elastic bands.</li>
		<li>For extended monitoring, under ultrasound guidance, place a three-lumen central venous catheter and a large-bore venous catheter (7 Fr.)&#160;into the internal jugular vein and a large-bore venous catheter (7 Fr.) for volume therapy. In addition, insert an arterial catheter into the internal carotid/cervical artery under ultrasound control for invasive blood pressure measurement (Figure 1B).</li>
		<li>Maintain anesthesia during organ retrieval via inhalation of isoflurane (0.8-1.5 vol%) and intravenous application of fentanyl (0.003-0.007 mg/kg body weight). Perform volume-controlled ventilation throughout the procedure. Apply 2,000 mg of sultamicillin for perioperative antibiosis and 250 mg of methylprednisolone intravenously.&#160;</li>
		<li>Administer a vasopressor such as norepinephrine intravenously to achieve a target mean arterial pressure of 60 mmHg. In addition, apply crystalloid solutions such as Ringer&#39;s lactate solution or colloid solutions such as fluid gelatins if necessary.</li>
		<li>Apply calcium gluconate (10%) and sodium bicarbonate (8.4%), glucose (40%), or potassium chloride (7.45%) intravenously with respect to blood gas analyses obtained every 30 min.</li>
	</ol>
	</li>
	<li>Recipient hepatectomy
	<ol>
		<li>Scrub the skin with antiseptic agent, e.g., povidone-iodine or isopropyl alcohol, and cover the animal with sterile drapes.</li>
		<li>Confirm an adequate depth of anesthesia by loss of the withdrawal response to toe pinch. Perform a midline laparotomy beginning at the xiphoid process by using monopolar cautery. Place an abdominal retractor and mobilize the intestine to the left of the donor. Cover the intestine with a moistened cloth.</li>
		<li>Place a suprapubic urinary catheter for the optimization of intraoperative volume management.</li>
		<li>Sever the falciform ligament and the triangular ligaments using scissors and bipolar cautery. After sufficient dissection of the liver, encircle both the suprahepatic and infrahepatic vena cavae inferior close to the liver parenchyma.</li>
		<li>Dissect and sever the common bile duct below the junction of the cystic duct between two ligatures (3-0 polyfilament).</li>
		<li>Incise the superficial peritoneal layer covering the hepatoduodenal ligament and identify the hepatic arteries shortly before entering the liver parenchyma. Dissect using bipolar cautery or the placement of clips, ligatures, or sutures.</li>
		<li>Dissect the abdominal aorta by incision in the midline (avascular layer) of the right and left diaphragmatic muscles. Prepare the aorta for the aortic anastomosis by removal of the surrounding tissue. 
		NOTE: This step only is required if an aortic anastomosis is performed. Otherwise, further dissect the hepatic artery/the hilar region to prepare for a conventional end-to-end anastomosis between the donor and recipient hepatic arteries.</li>
		<li>Perform recipient hepatectomy by placing an atraumatic vascular clamp on the portal vein, followed by atraumatic vascular clamps on the suprahepatic vena cava inferior (including the surrounding diaphragm while caudally retracting the liver) and the infrahepatic vena cava inferior.</li>
		<li>Sever all three vessels close to the liver parenchyma. Remove the recipient liver from the abdominal cavity. 
		NOTE: The clamping of the vessels marks the start of the anhepatic phase. During the anhepatic phase, the pigs are hemodynamically instable and require relevant amounts of vasopressors/catecholamines. The anesthesiologist should be prepared to apply norepinephrine and epinephrine. Keep the phase until reperfusion of the liver as short as possible. Communicate well with the anesthesiologist.</li>
	</ol>
	</li>
	<li>Donor liver engraftment
	<ol>
		<li>Place the donor liver into the abdominal cavity. Shorten the donor and/or recipient suprahepatic vena cava inferior to an adequate length while avoiding kinking or too much tension on the anastomosis.</li>
		<li>Place a single suture as a supporting thread (5-0 monofilament), adapting the right corner of the donor and recipient suprahepatic vena cavae inferior. Begin the dorsal side of the anastomosis from the left corner of the vessel(s) with a running suture (5-0 monofilament, double-armed).</li>
		<li>When reaching the right corner, remove the supporting thread, secure the running suture with a clamp, and continue with the ventral side of the anastomosis, again beginning from the left corner of the vessel(s). Tighten the suture with multiple knots without constricting the vessel diameter in order to avoid stenosis.</li>
		<li>Shorten the donor and/or recipient portal vein to an adequate length while avoiding kinking or too much tension on the anastomosis.</li>
		<li>Perform a vascular anastomosis of the donor and recipient portal vein analogous to steps 3.3.2-3.3.3 using a 6-0 monofilament, double-armed suture.</li>
		<li>Perform the porto-venous reperfusion by removing the vascular clamp, occluding the recipient portal vein, and occlude the donor infrahepatic vena cava inferior with a vascular clamp after draining approximately 200-400 mL of blood. Slowly remove the vascular clamp occluding the recipient suprahepatic vena cava inferior and search for active bleeding. 
		&#8203;NOTE: The removal of both clamps marks the end of the anhepatic phase. The amount of catecholamines required should significantly decrease shortly thereafter.</li>
		<li>Shorten the donor and/or recipient infrahepatic vena cava inferior. Perform a vascular anastomosis of the donor and recipient infrahepatic vena cavae inferior analogous to steps 3.3.2-3.3.3 using a 5-0 monofilament, double-armed suture. Remove the clamps occluding the donor and recipient infrahepatic vena cavae inferior.</li>
		<li>Prepare an elliptic aortal patch (Carrel patch) with a diameter of approximately 1-1.5, cm depending on anatomical circumstances, using scissors. Clamp the abdominal aorta with an atraumatic Cooley vascular clamp and make an incision by using a scalpel. Enlarge the incision using scissors to fit the patch.</li>
		<li>Begin the aortic anastomosis with a running suture (6-0 monofilament, double-armed) at the cranial corner of the incision/patch. When reaching the caudal corner, secure the running suture with a clamp and complete the anastomosis again beginning at the cranial corner. Tighten the suture with multiple knots and slowly remove the vascular clamp. 
		NOTE: Clamping of the abdominal aorta will significantly affect the blood pressure of the pig. Communicate well with the anesthesiologist.</li>
		<li>Place a hemostatic gauze around the arterial anastomosis. Place a catheter in the common bile duct and secure it with a single ligature. Make sure not to occlude the diameter of the catheter.</li>
		<li>Close the abdomen temporarily by adapting the muscular fascia and the skin with a running suture and cover the abdomen with cling film and/or drapes to avoid thermal loss. 
		NOTE: If the scientific objectives require a chronic model, perform an end-to-end anastomosis between the donor and recipient bile duct, close the abdomen with separate running sutures for the peritoneum and the muscular fascia, and close the skin with single sutures.</li>
		<li>At the end of follow-up, inject a lethal dose of 5,000 mg of pentobarbital sodium for intraoperative euthanasia.</li>
	</ol>
	</li>
</ol>

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J., 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=Hemodynamic+alterations+during+orthotopic+liver+experimental+transplantation+in+pigs.">Hemodynamic alterations during orthotopic liver experimental transplantation in pigs.</a> Acta Cirurgica Brasileria. 23 (2), 135-139 (2008).</li><li>Canedo, B. 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=Liver+autotransplantation+in+pigs+without+venovenous+bypass:+A+simplified+model+using+a+supraceliac+aorta+cross-clamping+maneuver.">Liver autotransplantation in pigs without venovenous bypass: A simplified model using a supraceliac aorta cross-clamping maneuver.</a> Annals of Transplantation. 20, 320-326 (2015).</li><li>Battersby, C., Hickman, R., Saunders, S. J., Terblanche, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Liver+function+in+the+pig.+1.+The+effects+of+30+minutes'+normothermic+ischaemia.">Liver function in the pig. 1. The effects of 30 minutes' normothermic ischaemia.</a> The British Journal of Surgery. 61 (1), 27-32 (1974).</li><li>Kaiser, G. M., Heuer, M. M., Fr&#252;hauf, N. R., K&#252;hne, C. A., Broelsch, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=General+handling+and+anesthesia+for+experimental+surgery+in+pigs.">General handling and anesthesia for experimental surgery in pigs.</a> Journal of Surgical Research. 130 (1), 73-79 (2006).</li><li>Oike, 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=Simplified+technique+of+orthotopic+liver+transplantation+in+pigs.">Simplified technique of orthotopic liver transplantation in pigs.</a> Transplantation. 71 (2), 328-331 (2001).</li><li>Heuer, 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=Liver+transplantation+in+swine+without+venovenous+bypass.">Liver transplantation in swine without venovenous bypass.</a> European Surgical Research. 45 (1), 20-25 (2010).</li><li>Fondevila, 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=Step-by-step+guide+for+a+simplified+model+of+porcine+orthotopic+liver+transplant.">Step-by-step guide for a simplified model of porcine orthotopic liver transplant.</a> The Journal of Surgical Research. 167 (1), 39-45 (2011).</li></ol>

The authors have nothing to disclose.

Recent technical developments such as the introduction of machine perfusion have the potential to revolutionize the field of liver transplantation. In order to translate graft reconditioning or modification concepts into clinical settings, reproducible transplant models in large animals are inevitable.
After the initial introduction of porcine orthotopic liver transplantation, several authors have worked on the improvement of these techniques over the past five&#160;decades. Differences within...

Liver transplantation remains to be the only chance for survival in a variety of different diseases leading to acute or chronic hepatic failure. Since its first successful application in mankind in 1963 by Thomas E. Starzl, the concept of liver transplantation has evolved into a reliable treatment option applied worldwide, mainly as a result of advancements in the understanding of the immune system, the development of modern immunosuppression, and the optimization of perioperative care and surgical techniques1,2. However, aging populations and a higher demand for organs have resulted in donor shortages, with increased use of marginal grafts from extended criteria donors and the emergence of new challenges in the past decades. The introduction and widespread implementation of organ machine perfusion is believed to open up an array of possibilities with regard to graft reconditioning and modulation and to help mitigate organ shortages and reduce waiting list mortality3,4,5,6.
In order to evaluate these concepts and their effects in vivo, translational transplant models are necessary7. In 1983, Kamada et al. introduced an efficient orthotopic liver transplant model in rats that has since been extensively modified and applied by workgroups around the globe8,9,10,11. The orthotopic liver transplant model in mice is technically more demanding, but also more valuable in terms of immunological transferability, and was first reported in 1991 by Qian et al.12. Despite advantages regarding availability, animal welfare, and costs, rodent models are limited in their applicability in clinical settings7. Hence, large animal models are required.
In recent years, pigs have become the main animal species used for translational research due to their anatomical and physiological similarities with humans. Furthermore, current progress in the field of xenotransplantation might further increase the importance of pigs as research objects13,14.
Garnier et al. described a liver transplant model in pigs as early as 196515. Several authors, including Calne et al. in 1967 and Chalstrey et al. in 1971, subsequently reported modifications, ultimately leading to a safe and feasible concept of experimental porcine liver transplantation in the decades to follow16,17,18,19,20,21.
More recently, different work groups have provided data with regard to current issues in liver transplantation using a technique of porcine orthotopic liver transplantation, almost invariably including an active or passive veno-venous, i.e., porto-caval, bypass19,22. The reason for this is a species-specific intolerance to the clamping of the vena cava inferior and the portal vein during the anhepatic phase due to a comparatively larger intestine and fewer porto-caval or cavo-caval shunts (e.g., lack of a vena azygos), resulting in increased perioperative morbidity and mortality23. Vena cava inferior-sparing transplant techniques applied in human recipients as an alternative are not feasible as the porcine vena cava inferior is encased by hepatic tissue23.
However, the usage of a veno-venous bypass further increases technical and logistical complexity in an already demanding surgical procedure, therefore possibly preventing workgroups from attempting implementation of the model altogether. Apart from the direct physiological and immunological effects of a bypass, some authors have pointed out the significant morbidity such as blood loss or air embolism during shunt placement and the need for a simultaneous splenectomy, potentially affecting short- and long-term results after engraftment24,25.
The following protocol describes a simple technique of porcine orthotopic liver transplantation after static cold storage of donor organs for 20 h, representing extended criteria donor conditions without the usage of a veno-venous bypass during engraftment, including donor liver procurement, back-table preparation, recipient hepatectomy, and anesthesiological pre- and intraoperative management.
This model should be of special interest for surgical workgroups focusing on the immediate postoperative course, ischemia-reperfusion injury, the reconditioning of extended criteria donor organs, and associated immunological mechanisms.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Abdominal retractor</td>
			<td>No Company Name available</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Aortic clamp, straight</td>
			<td>Firma Martin</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Arterial Blood Sampler Aspirator (safePICOAspirator) 1.5 mL</td>
			<td>Radiometer Medical ApS</td>
			<td>956-622</td>
			<td></td>
		</tr>
		<tr>
			<td>Atropine (Atropinsulfat 0.5 mg/1 mL)</td>
			<td>B.Braun</td>
			<td>648037</td>
			<td></td>
		</tr>
		<tr>
			<td>Backhaus clamp</td>
			<td>Bernshausen</td>
			<td>BF432</td>
			<td></td>
		</tr>
		<tr>
			<td>Bipolar forceps, 23 cm&#160;</td>
			<td>SUTTER</td>
			<td>780222 SG</td>
			<td></td>
		</tr>
		<tr>
			<td>Bowl 5 L, 6 L, 9 L</td>
			<td>Chiru-Instrumente</td>
			<td>35-114327</td>
			<td></td>
		</tr>
		<tr>
			<td>Braunol Braunoderm</td>
			<td>B.Braun</td>
			<td>3881059</td>
			<td></td>
		</tr>
		<tr>
			<td>Bulldog clamp</td>
			<td>Aesculap</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Button canula</td>
			<td>Krauth + Timmermann GmbH</td>
			<td>1464LL1B</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium gluconate (2.25 mmol/10 mL (10%))</td>
			<td>B.Braun</td>
			<td>2353745</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell Saver (Autotransfusion Reservoir)</td>
			<td>Fresenius Kabi AG</td>
			<td>9108471</td>
			<td></td>
		</tr>
		<tr>
			<td>Central venous catheter 7Fr., 3 Lumina, 30 cm 0.81 mm</td>
			<td>Arrow</td>
			<td>AD-24703</td>
			<td></td>
		</tr>
		<tr>
			<td>Clamp</td>
			<td>INOX</td>
			<td>B-17845&#160; /&#160; BH110&#160; / B-481</td>
			<td></td>
		</tr>
		<tr>
			<td>Clamp</td>
			<td>Aesculap</td>
			<td>AN909R</td>
			<td></td>
		</tr>
		<tr>
			<td>Clamp, 260 mm</td>
			<td>Fehling Instruments GMbH &#38;Co.KG</td>
			<td>ZAU-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Clip Forceps, medium</td>
			<td>Ethicon</td>
			<td>LC207</td>
			<td></td>
		</tr>
		<tr>
			<td>Clip forceps, small</td>
			<td>Ethicon&#160;</td>
			<td>LC107</td>
			<td></td>
		</tr>
		<tr>
			<td>CPDA-1 solution</td>
			<td>Fresenius Kabi AG</td>
			<td>41SD09AA00</td>
			<td></td>
		</tr>
		<tr>
			<td>Custodiol (Histidin-Tryptophan-Ketogluterat-Solution)</td>
			<td>Dr.Franz K&#246;hler Chemie GmbH</td>
			<td>2125921</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting scissors</td>
			<td>LAWTON&#160; 05-0641&#160;</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting scissors, 180 mm</td>
			<td>Metzenbaum&#160;</td>
			<td>BC606R</td>
			<td></td>
		</tr>
		<tr>
			<td>Endotracheal tube 8.0 mm</td>
			<td>Covetrus</td>
			<td>800764</td>
			<td></td>
		</tr>
		<tr>
			<td>Epinephrine (Adrenalin 1:1000)</td>
			<td>InfectoPharm</td>
			<td>9508734</td>
			<td></td>
		</tr>
		<tr>
			<td>Falcon Tubes 50ml</td>
			<td>Greiner&#160;</td>
			<td>227 261 L</td>
			<td></td>
		</tr>
		<tr>
			<td>Femoralis clamp</td>
			<td>Ulrich&#160;</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Fentanyl 0.1mg</td>
			<td>PanPharma</td>
			<td>00483</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps, anatomical</td>
			<td>Martin</td>
			<td>12-100-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps, anatomical, 250 mm</td>
			<td>Aesculap</td>
			<td>BD052R</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps, anatomical, 250 mm</td>
			<td>Aesculap</td>
			<td>BD032R</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps, anatomical, 250 mm&#160;</td>
			<td>Aesculap</td>
			<td>BD240R</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps, surgical</td>
			<td>Bernshausen</td>
			<td>BD 671</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps, surgical</td>
			<td>INOX</td>
			<td>B-1357</td>
			<td></td>
		</tr>
		<tr>
			<td>G40 solution</td>
			<td>Serag Wiessner</td>
			<td>10755AAF</td>
			<td></td>
		</tr>
		<tr>
			<td>Gelafundin ISO solution 40 mg/mL</td>
			<td>B. Braun</td>
			<td>210257641</td>
			<td></td>
		</tr>
		<tr>
			<td>Guidewire with marker</td>
			<td>Arrow</td>
			<td>14F21E0236</td>
			<td></td>
		</tr>
		<tr>
			<td>Haemostatic gauze (&#34;Tabotamp&#34;&#160; 5 x 7.5 cm)</td>
			<td>Ethicon</td>
			<td>474273</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin sodium 25,000IE</td>
			<td>Ratiopharm</td>
			<td>W08208A</td>
			<td></td>
		</tr>
		<tr>
			<td>Hico-Aquatherm 60</td>
			<td>Hospitalwerk</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Infusion Set Intrafix</td>
			<td>B.Braun</td>
			<td>4062981 L</td>
			<td></td>
		</tr>
		<tr>
			<td>Intrafix SafeSet 180 cm</td>
			<td>B.Braun</td>
			<td>4063000</td>
			<td></td>
		</tr>
		<tr>
			<td>Introcan Safety, 18 G&#160;</td>
			<td>B.Braun</td>
			<td>4251679-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Isofluran CP</td>
			<td>CP-Pharma</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Large-bore venous catheter, 7Fr.</td>
			<td>Edwards Lifesciences</td>
			<td>I301F7</td>
			<td></td>
		</tr>
		<tr>
			<td>Ligaclip, medium</td>
			<td>Ethicon</td>
			<td>LT200</td>
			<td></td>
		</tr>
		<tr>
			<td>Ligaclip, small</td>
			<td>Ethicon&#160;</td>
			<td>LT100</td>
			<td></td>
		</tr>
		<tr>
			<td>Material scissors</td>
			<td>Martin&#160;</td>
			<td>11-285-23</td>
			<td></td>
		</tr>
		<tr>
			<td>Methylprednisolone (Urbason solubile forte 250 mg)</td>
			<td>Sanofi</td>
			<td>7823704</td>
			<td></td>
		</tr>
		<tr>
			<td>Monopolar ERBE ICC 300</td>
			<td>Fa. Erbe</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl solution (0.9%)</td>
			<td>Baxter</td>
			<td>1533</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle holder</td>
			<td>Aesculap</td>
			<td>BM36</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle holder</td>
			<td>Aesculap</td>
			<td>BM035R</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle holder</td>
			<td>Aesculap</td>
			<td>BM 67</td>
			<td></td>
		</tr>
		<tr>
			<td>Neutral electrode</td>
			<td>Erbe Elektromedizin GmbH T&#252;bingen</td>
			<td>21191 - 060</td>
			<td></td>
		</tr>
		<tr>
			<td>Norepinephrine (Sinora)</td>
			<td>Sintetica GmbH</td>
			<td>04150124745717</td>
			<td></td>
		</tr>
		<tr>
			<td>Omniflush Sterile Filed 10 mL</td>
			<td>B.Braun</td>
			<td>3133335</td>
			<td></td>
		</tr>
		<tr>
			<td>Original Perfusorline 300 cm</td>
			<td>B.Braun</td>
			<td>21E26E8SM3</td>
			<td></td>
		</tr>
		<tr>
			<td>Overhold clamp</td>
			<td>INOX</td>
			<td>BH 959</td>
			<td></td>
		</tr>
		<tr>
			<td>Overhold clamp</td>
			<td>Ulrich</td>
			<td>CL 2911</td>
			<td></td>
		</tr>
		<tr>
			<td>Pentobarbital sodium(Release 500 mg/mL)</td>
			<td>WDT, Garbsen</td>
			<td>21217</td>
			<td></td>
		</tr>
		<tr>
			<td>Perfusers</td>
			<td>B.Braun</td>
			<td>49-020-031</td>
			<td></td>
		</tr>
		<tr>
			<td>Perfusor Syringe 50 mL</td>
			<td>B.Braun</td>
			<td>8728810F</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dishes&#160; 92 x 17 mm</td>
			<td>Nunc</td>
			<td>150350</td>
			<td></td>
		</tr>
		<tr>
			<td>Poole Suction Instrument Argyle flexibel</td>
			<td>Covidien, Mansfield USA</td>
			<td>20C150FHX</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium chloride (7.45%)</td>
			<td>B.Braun</td>
			<td>4030539078276</td>
			<td></td>
		</tr>
		<tr>
			<td>Pressure measurement set</td>
			<td>Codan pvb Medical GmbH</td>
			<td>957179</td>
			<td></td>
		</tr>
		<tr>
			<td>Propofol (1%)</td>
			<td>CP-Pharma</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>S-Monovette 2.6 mL K3E</td>
			<td>Sarstedt</td>
			<td>04.1901</td>
			<td></td>
		</tr>
		<tr>
			<td>S-Monovette 2.9 mL 9NC</td>
			<td>Sarstedt</td>
			<td>04.1902</td>
			<td></td>
		</tr>
		<tr>
			<td>S-Monovette 7.5 mL Z-Gel</td>
			<td>Sarstedt</td>
			<td>11602</td>
			<td></td>
		</tr>
		<tr>
			<td>Sartinski clamp</td>
			<td>Aesculap</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel&#160; No.11</td>
			<td>Feather Safety Razor Co.LTD</td>
			<td>02.001.40.011</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>INOX&#160;</td>
			<td>BC 746</td>
			<td></td>
		</tr>
		<tr>
			<td>Seldinger Arterial catheter</td>
			<td>Arrow</td>
			<td>SAC-00520</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium bicarbonate (8.4%)</td>
			<td>B.Braun</td>
			<td>212768082</td>
			<td></td>
		</tr>
		<tr>
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			<td>B.Braun</td>
			<td>4899719</td>
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			<td>Sterofundin ISO solution</td>
			<td>B.Braun</td>
			<td>No Catalog Number available</td>
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			<td>Dahlhausen</td>
			<td>07.068.25.301</td>
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			<td>Aesculap</td>
			<td>No Catalog Number available</td>
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			<td>ConvaTec</td>
			<td>5365049</td>
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			<td>Pfizer</td>
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			<td>UK&#160; 1F02772</td>
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			<td>31654</td>
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			<td>Ethicon</td>
			<td>VCP 1218 H</td>
			<td></td>
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			<td>Suture Vicryl 4-0</td>
			<td>Ethicon</td>
			<td>V392H</td>
			<td></td>
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			<td>Suture, Prolene 4-0</td>
			<td>Ethicon</td>
			<td>7588 H</td>
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			<td>Suture, Prolene 5-0, double armed</td>
			<td>Ethicon&#160;</td>
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			<td>Suture, Prolene 5-0, single armed</td>
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			<td>Suture, Prolene: blau 3-0&#160;</td>
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			<td>Serag Wiessner</td>
			<td>353784</td>
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			<td>Syringe 2 mL, 5 mL, 10 mL, 20 mL</td>
			<td>B.Braun</td>
			<td>4606027V</td>
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			<td>TransferSet &#34;1D/X-double&#34; steril 330 cm</td>
			<td>Fresenius Kabi AG</td>
			<td>2877101</td>
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		<tr>
			<td>Ultrasound Butterfly IQ+</td>
			<td>Butterfly Network Inc.</td>
			<td>850-20014</td>
			<td></td>
		</tr>
		<tr>
			<td>Ventilator &#34;Oxylog Dr&#228;ger Fl&#34;</td>
			<td>Dr&#228;ger Medical AG</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Yankauer Suction</td>
			<td>Medline</td>
			<td>RA19GMD</td>
			<td></td>
		</tr>
		<tr>
			<td>Zoletil 100 mg/mL&#160; (50 mg Zolazepam, 50 mg tiletamin)</td>
			<td>Virbac</td>
			<td>794-861794861</td>
			<td></td>
		</tr>
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porcine liver transplantation without veno-venous bypass as an extended criteria donor model

Liver transplantation is regarded as the gold standard for the treatment of a variety of fatal hepatic diseases. However, unsolved issues of chronic graft failure, ongoing organ donor shortages, and the increased use of marginal grafts call for the improvement of current concepts, such as the implementation of organ machine perfusion. In order to evaluate new methods of graft reconditioning and modulation, translational models are required. With respect to anatomical and physiological similarities to humans and recent progress in the field of xenotransplantation, pigs have become the main large animal species used in transplantation models. After the initial introduction of a porcine orthotopic liver transplant model by Garnier et al. in 1965, several modifications have been published over the past 60 years.
Due to specifies-specific anatomical traits, a veno-venous bypass during the anhepatic phase is regarded as a necessity to reduce intestinal congestion and ischemia resulting in hemodynamic instability and perioperative mortality. However, the implementation of a bypass increases the technical and logistical complexity of the procedure. Furthermore, associated complications such as air embolism, hemorrhage, and the need for a simultaneous splenectomy have been reported previously.
In this protocol, we describe a model of porcine orthotopic liver transplantation without the use of a veno-venous bypass. The engraftment of donor livers after static cold storage of 20 h - simulating extended criteria donor conditions - demonstrates that this simplified approach can be performed without significant hemodynamic alterations or intraoperative mortality and with regular uptake of liver function (as defined by bile production and liver-specific CYP1A2 metabolism). The success of this approach is ensured by an optimized surgical technique and a sophisticated anesthesiologic volume and vasopressor management.
This model should be of special interest for workgroups focusing on the immediate postoperative course, ischemia-reperfusion injury, associated immunological mechanisms, and the reconditioning of extended criteria donor organs.

The technique presented in this protocol has provided reliable and reproducible results in terms of hemodynamic stability and animal survival throughout the procedure, as well as graft function in the postoperative course.
Most recently, we applied the model for the study of ischemia-reperfusion injury and therapeutic interventions mitigating detrimental effects in the immediate postoperative course. Upon retrieval and 20 h of static cold storage, liver grafts (with a mean weight of 983.38 g) ...

Watch this Scientific Journal Video about Porcine Liver Transplantation Without Veno-Venous Bypass As an Extended Criteria Donor Model at JoVE.com

Porcine Liver Transplantation Without Veno-Venous Bypass As an Extended Criteria Donor Model

In this protocol, a model of porcine orthotopic liver transplantation after static cold storage of donor organs for 20 h without the use of a veno-venous bypass during engraftment is described. The approach uses a simplified surgical technique with minimization of the anhepatic phase and sophisticated volume and vasopressor management.

Liver transplantation is regarded as the gold standard for the treatment of a variety of fatal hepatic diseases. However, unsolved issues of chronic graft ...

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2.2K Views.  Hannover Medical School. In this protocol, a model of porcine orthotopic liver transplantation after static cold storage of donor organs for 20 h without the use of a veno-venous bypass during engraftment is described. The approach uses a simplified surgical technique with minimization of the anhepatic phase and sophisticated volume and vasopressor management.

Video: Porcine Liver Transplantation Without Veno-Venous Bypass As an Extended Criteria Donor Model

Orthotopic Liver Transplantation in Rats

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

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

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

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

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

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

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

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

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

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

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

Ex Situ Normothermic Machine Perfusion of Donor Livers

In contrast to conventional static cold preservation (0-4 &#176;C), ex situ machine perfusion may provide better preservation of donor livers. Continuous perfusion of organs provides the opportunity to improve organ quality and allows ex situ viability assessment of donor livers prior to transplantation. This video article provides a step by step protocol for ex situ normothermic machine perfusion (37 &#176;C) of human donor livers using a device that provides a pressure and temperature controlled pulsatile perfusion of the hepatic artery and continuous perfusion of the portal vein. The perfusion fluid is oxygenated by two hollow fiber membrane oxygenators and the temperature can be regulated between 10 &#176;C and 37 &#176;C. During perfusion, the metabolic activity of the liver as well as the degree of injury can be assessed by biochemical analysis of samples taken from the perfusion fluid. Machine perfusion is a very promising tool to increase the number of livers that are suitable for transplantation.

Ex Situ Normothermic Machine Perfusion of Donor Livers

Here we present a protocol describing oxygenated ex situ machine perfusion of donor liver grafts. This article contains a step by step protocol to procure and prepare the liver graft for machine perfusion, prepare the perfusion fluid, prime the perfusion machine and perform oxygenated normothermic machine perfusion of the liver graft.

In contrast to conventional static cold preservation (0-4 &#176;C), ex situ machine perfusion may provide better preservation of donor livers. Continuous ...

Cardiopulmonary Bypass in a Mouse Model: A Novel Approach

As prolonged cardiopulmonary bypass becomes more essential during cardiac interventions, an increasing clinical demand arises for procedure optimization and for minimizing organ damage resulting from prolonged extracorporal circulation. The goal of this paper was to demonstrate a fully functional and clinically relevant model of cardiopulmonary bypass in a mouse. We report on the device design, perfusion circuit optimization, and microsurgical techniques. This model is an acute model, which is not compatible with survival due to the need for multiple blood drawings. Because of the range of tools available for mice (e.g., markers, knockouts, etc.), this model will facilitate investigation into the molecular mechanisms of organ damage and the effect of cardiopulmonary bypass in relation to other comorbidities.

This paper describes how to perform cardiopulmonary bypass in mice. This novel model will facilitate the investigation of the molecular mechanisms involved in organ damage.

As prolonged cardiopulmonary bypass becomes more essential during cardiac interventions, an increasing clinical demand arises for procedure optimization ...

Re-Arterialized Rat Partial Liver Transplantation with an in vivo Vessel-Oriented 70% Hepatectomy

Split liver transplantation and living liver donor liver transplantation were developed in the clinic to utilize liver organs in a more efficient manner. To better understand the mechanism behind these surgical procedures, a rat partial liver transplantation (PLTx) model was established for relevant surgical studies. Because of the complexity of the rat PLTx model, a protocol with detailed descriptions is required. An article published previously reported a protocol in which ex vivo hepatectomy was used to achieve 50% rat PLTx. In contrast to this protocol, we introduced a re-arterialized PLTx with an in vivo 70% hepatectomy. An updated vessel-oriented hepatectomy was incorporated into the rat PLTx to refine the microsurgical procedure. The portal veins and hepatic arteries of the left lateral lobe and the median lobe were individually dissected and ligated before removal of the liver parenchyma, thereby decreasing the probability of bleeding in the remnant liver stump. Furthermore, an end-to-side vessel anastomosis between the common hepatic artery and the enlarged proper hepatic artery was introduced to re-arterialize the hepatic artery. By using this end-to-side vessel anastomosis technique, the diameter of the anastomosis was enlarged, thereby decreasing the difficulty of hand suture and maintaining a high rate of anastomotic patency. Moreover, the cuff anastomosis of the infrahepatic vena cava was slightly modified. A section of circumferential liver parenchyma around the vena cava of a recipient was preserved during cuff anastomosis to maintain the three-dimensional shape of the vascular lumen. This section of liver parenchyma was removed after completing the anastomosis. With this modification, the step involving placement of stay sutures was omitted, thereby further shortening the cuff anastomosis time. By using this protocol of rat PLTx, a low liver enzyme level, an intact liver lobular architecture and a high survival rate were achieved after microsurgery.

Re-Arterialized Rat Partial Liver Transplantation with an in vivo Vessel-Oriented 70% Hepatectomy

Here, a protocol involving re-arterialized rat partial liver transplantation is presented. Specifically, 70% liver was resected in vivo by using an updated technique of vessel-oriented hepatectomy. The hepatic artery was reconstructed in an end-to-side manner. The cuff technique was modified to shorten the anastomosis time of the infrahepatic vena cava.

Split liver transplantation and living liver donor liver transplantation were developed in the clinic to utilize liver organs in a more efficient manner. ...

Veno-Venous Extracorporeal Membrane Oxygenation in a Mouse

The use of extracorporeal membrane oxygenation (ECMO) has increased substantially in recent years. ECMO has become a reliable and effective therapy for acute as well as end-stage lung diseases. With the increase in clinical demand and prolonged use of ECMO, procedural optimization and prevention of multi-organ damage are of critical importance. The aim of this protocol is to present a detailed technique of veno-venous ECMO in a non-intubated, spontaneously breathing mouse. This protocol demonstrates the technical design of the ECMO and surgical steps. This murine ECMO model will facilitate the study of pathophysiology related to ECMO (e.g., inflammation,bleeding and thromboembolic events). Due to the abundance of genetically modified mice, the molecular mechanisms involved in ECMO-related complications can also be dissected.

Here we present a protocol describing the technique of veno-venous extracorporeal membrane oxygenation (ECMO) in a non-intubated, spontaneously breathing mouse. This murine model of ECMO can be effectively implemented in experimental studies of acute and end-stage lung diseases.

The use of extracorporeal membrane oxygenation (ECMO) has increased substantially in recent years. ECMO has become a reliable and effective therapy for ...

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 ...

Murine Precision-Cut Liver Slices as an Ex Vivo Model of Liver Biology

Understanding the mechanisms of liver injury, hepatic fibrosis, and cirrhosis that underlie chronic liver diseases (i.e., viral hepatitis, non-alcoholic fatty liver disease, metabolic liver disease, and liver cancer) requires experimental manipulation of animal models and in vitro cell cultures. Both techniques have limitations, such as the requirement of large numbers of animals for in vivo manipulation. However, in vitro cell cultures do not reproduce the structure and function of the multicellular hepatic environment. The use of precision-cut liver slices is a technique in which uniform slices of viable mouse liver are maintained in laboratory tissue culture for experimental manipulation. This technique occupies an experimental niche that exists between animal studies and in vitro cell culture methods. The presented protocol describes a straightforward and reliable method to isolate and culture precision-cut liver slices from mice. As an application of this technique, ex vivo liver slices are treated with bile acids to simulate cholestatic liver injury and ultimately assess the mechanisms of hepatic fibrogenesis.

This protocol provides a simple and reliable method for the production of viable precision-cut liver slices from mice. The ex vivo tissue samples can be maintained under laboratory tissue culture conditions for multiple days, providing a flexible model to examine liver pathobiology.

Understanding the mechanisms of liver injury, hepatic fibrosis, and cirrhosis that underlie chronic liver diseases (i.e., viral hepatitis, non-alcoholic ...

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.

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. ...