The study was supported by research grants of the Roche Organ Transplant Research Foundation (ROTRF) and Astellas. Markus Selzner was supported by an ASTS Career Development Award. Matthias Knaak was supported by the Astellas Research Scholarship. We thank Uwe Mummenhoff and the Birmingham family for their generous support.

A schematic overview of the protocol is presented in Figure 1.
<img alt="Figure 1" src="/files/ftp_upload/51419/51419fig1highres.jpg" /> 
Figure 1. Study protocol. The porcine study design of liver injury is based on a donation after cardiac death (DCD) model. After dissection of all liver vessels, cardiac death is induced followed by 45 min of warm graft ischemia. To simulate a graft transport in between the donor and recipient hospitals in a clinical setting, the graft is stored on ice for 4 hr after cold, dual flush. After cold storage, the organ is subnormothermic perfused for 6 hr in order to assess the perfusion stability. In a transplant model, the perfusion time could be shorter in order to recharge energy storage and to assess the organ viability. <a href="https://www.jove.com/files/ftp_upload/51419/51419fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
1. Animals
NOTE: Male Yorkshire pigs, 30-35 kg, were utilized for this study. All animals received humane care in compliance with the &#8216;&#8216;Principles of Laboratory Animal Care&#8217;&#8217; formulated by the National Society for Medical Research and the &#8216;&#8216;Guide for the Care of Laboratory Animals&#8217;&#8217; published by the National Institutes of Health. The Animal Care Committee of the Toronto General Research Institute approved all studies.
2. Organ Retrieval
<ol>
	<li>House male Yorkshire pigs in research facilities for 1 week before perfusion/transplantation to reduce the level of stress and to accustom the animals to the housing conditions. Less than 2&#160;days of housing inside the facility will lead to a stress-induced physical reaction, which can alter the perfusion&#8217;s outcome22,23.</li>
	<li>Anesthetize pigs by an intramuscular (i.m.) injection of a mixture of ketamine (25 mg/kg), atropine (0.04 mg/kg), and midazolam (0.15 mg/kg).</li>
	<li>Prior to intubation, ensure the pig breathes spontaneously 2 L of O2 dosed with 5% isoflurane. Spray the vocal chords with 2% lidocaine 2 min before intubation to avoid vocal cord spasms. For example, for a 35 kg pig use a 6.5 fr. tracheal tube. Block the tracheal tube with 3-5 ml of room air.</li>
	<li>After intubation, use capnometry to confirm correct intubation. Lower the isoflurane gas to 2%. Set the ventilator to 14-16 breaths/min and a tidal volume of 10 ml/kg bodyweight.</li>
	<li>Place a 20 G intravenous (i.v.) catheter in one of the ear veins to allow infusion of Ringer&#8217;s lactate solution (200 ml per hr). Then scrub the pig and cover it with sterile drapes.</li>
	<li>Make a midline incision followed by a left lateral extension. Use a towel to cover large and small bowels and move them to the left side.</li>
	<li>Separate inferior vena cava (IVC) and distal aorta from each other; ligate aorta branches to the back; isolate and free renal arteries from adherent tissue.</li>
	<li>Divide the falciforme ligament and the triangular ligament using cautery.</li>
	<li>Release the portal vein by an incision of the peritoneum between pancreas and portal vein. Tie off veins draining from the pancreas to the portal vein.</li>
	<li>Dissect the coeliac trunk below the portal vein and follow it backwards to the aorta. Surround the mesenteric artery with a 2-0 tie; surround the splenic and left gastric arteries, which branch posteriorly to the coeliac trunk. Dissect the coeliac trunk off the portal vein.</li>
	<li>Ligate the lymph vessels within the hepatoduodenal ligament to prevent lymphatic leakage. Divide the right gastric artery between ties. Ligate smaller veins. Separate the bile duct from the ligament and divide it distally after ligation.</li>
	<li>Dissect the aorta behind the diaphragm between heart and coeliac trunk; place a 2-0 tie around the aorta.</li>
	<li>Release the liver from the lower cava on the right side using electro cautery; use scissors for the upper part between cava and liver.</li>
	<li>Remove the gallbladder and cauterize bleeders from gallbladder bed.</li>
	<li>Administer i.v. 1,000 iU/kg donor weight of Heparin. For a DCD model, induce cardiac arrest by intracardial injection of 40 mval KCl 3 min after Heparin administration. Set cardiac arrest as the starting point of warm ischemia.</li>
	<li>For the perfusion, collect 1.6 L of pig blood in CPDA bags (citrate, phosphate, dextrose, adenosine) immediately after cardiac death. Perform a soft spin (2,000 x g without brake). Remove the plasma and the buffy coat under sterile condition (biosafety cabinet class II) and store the erythrocytes in CPDA bags for transfusion.</li>
	<li>Cannulate portal vein and aorta with organ flush lines. Tie off the previously set ties around femoral, renal, splenic, mesenteric, and left gastric arteries as well as the upper aorta. For a heart beating donor (HBD) model, perform the cannulation of aorta and portal vein under heart beating conditions.</li>
	<li>After 45 min warm ischemia, flush the liver with University of Wisconsin (UW) solution using dual perfusion via aorta (pressure bag) and portal vein (gravity driven).</li>
	<li>Cut the liver out of the pig, leaving all remaining vessels long.</li>
	<li>During back-table preparation, clamp the upper IVC using a Satinsky clamp and flush the liver a second time with about 0.5 L of UW solution retrogradely via lower IVC until the portal vein outflow is clear.</li>
	<li>Tie off all arterial branches from the aorta and coeliac trunk. Perform an arterial back-table pressure perfusion with about 0.5 L of UW solution.</li>
	<li>Flush the bile duct using UW solution.</li>
	<li>Cannulate the upper and lower parts of the IVC using 1/2&#8221; x 3/8&#8221; reducers with Luer Lock; cannulate the portal vein and liver artery using 3/8&#8221; x 1/4&#8221; and 1/4&#8221; x 3/8&#8221; reducers with Luer Lock. Use the upper and lower vena cava as venous drainage.</li>
	<li>Place the liver in an organ bag, close the organ bag, and store the liver on ice until the perfusion has started.</li>
</ol>
3. Ex vivo Liver Perfusion
<ol>
	<li>Prepare the perfusion solution containing 2,000 ml Steen solution, 400 ml washed erythrocytes, 550 mg sodium pyruvate, 100 ml amino acid solution (10% Travasol), 10 mg calcium gluconate, 1,000 IE rapid acting insulin, 1 g cefazolin, 500 mg metronidazol, and 10,000 iU heparin. Add other molecules for vasodilatation, immunosuppression, scavenging of reactive oxygen species, or liver cell treatment based on the particular study protocol.</li>
	<li>For the dialysis component, use standard dialysate containing 3.5 mM potassium, 25 mM bicarbonate, 27 mM glucose, as well as 275 mg/L pyruvate.</li>
	<li>Set up the perfusion circuit (schema see Figure 2). 
	<img alt="Figure 2" src="/files/ftp_upload/51419/51419fig2highres.jpg" /> 
	Figure 2. Circuit set up. Coming from the main reservoir as the starting and end point, the perfusion solution is driven by a centrifugal pump through an oxygenator. Right after oxygenation of the solution, the circuit splits into a smaller line running to a dialyzer unit for electrolyte homeostasis and a bigger line running to a leukocyte filter for reduction of remnant white blood cells (WBC). The solution that runs through the dialyzer returns to the main reservoir. After the leukocyte filter, the circuit splits up again into 2 equal lines. One line runs at high pressure (around 60 mmHg) directly into the aorta to perfuse the hepatic artery. The other line drains into a second reservoir. From this reservoir the portal vein is perfused. The pressure of the portal perfusion depends on the elevation energy of the reservoir&#8217;s solution level (around 2-6 mmHg). All fluids are drained via the infra- and supra-hepatic cava back into the main reservoir. For gravity reduction and homogeneous perfusion, the liver is placed in a pool filled with temperature regulated water. It is separated from the water by an impermeable membrane and it swims in a perfusion suspension. <a href="https://www.jove.com/files/ftp_upload/51419/51419fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
	<ol>
		<li>Collect all fluids from the circuit in a 3 L reservoir (main reservoir) and clamp the outflow.</li>
		<li>Connect the outflow to a centrifugal pump followed by a commercial oxygenator.</li>
		<li>Behind the oxygenator, split up the tubing into 2 lines. Connect one line to a dialyzer and drain it back into the main reservoir. Connect the second line to a leukocyte reduction filter.</li>
		<li>Split up the line after the leukocyte reduction filter into an arterial line, which supplies perfusion solution to the hepatic artery, and a portal venous line delivering perfusion solution to a second reservoir, which drains into the portal vein by gravity outflow. Clamp the portal inflow.</li>
		<li>Connect the arterial line to a vena cava line draining into the main reservoir for fluid recollection.</li>
		<li>For collection of ascites or leakage of perfusion solution from the liver, prepare a suction line connected to the main reservoir.</li>
	</ol>
	</li>
	<li>Release the outflow clamp from the main reservoir and fill the circuit with the perfusion solution. Start the centrifugal pump at 1,500 rounds/min. The perfusion solution will drive through the arterial line into the vena cava line back into the main reservoir. Make sure all air is driven out the circuit.</li>
	<li>Turn on the gas supply to the oxygenator.</li>
	<li>Take the liver off the ice. Flush out the UW solution using saline.</li>
	<li>Place the liver, with its convex side down to facilitate the access to the vessels, ideally in a gravity-free environment to avoid organ compression at the contact surface. Use a heat- and coolable water bath. Set the starting temperature of the circuit and water bath to 20 &#176;C. Cover the water bath with an impermeable membrane and place the liver onto that membrane. Reduce gravity driven compression by submerging the liver with perfusion solution.</li>
	<li>Reduce the speed of the centrifugal pump to 1,000 rounds/min and place two clamps at the connection of arterial and vena cava lines. Then, cut the tubing in between the clamps. Using a 3-way connector, join both cava outflows and connect them to the vena cava line.</li>
	<li>Release the clamp from the arterial line, pour perfusion solution into the arterial cannula to get rid of bubbles, and connect the line to the cannula. Increase the centrifugal pump to 1,500 rounds/min. Release the second clamp from the vena cava line.</li>
	<li>Release the clamp from the portal venous reservoir in order to fill it up. Let perfusion solution pour into the portal cannula and connect it. Take special care of stable fluid levels in the portal reservoir.</li>
	<li>Connect pressure lines to the Luer Locks of the arterial, portal, and vena cava cannulas.</li>
	<li>To mimic physiological conditions, apply treatments into the right vessel. Inject glucose into the portal vein and not into the arterial line in order to establish a gradient mimicking an increased portal venous glucose gradient and inducing glycogen synthesis24,25.</li>
	<li>After connecting the liver to the circuit, raise the temperature to 33 &#176;C within 60 min.</li>
	<li>Aim for an arterial starting flow at about 250 ml/min at 40 mmHg. This may reach 700 ml/min during perfusion once the pressure is increased up to 70 mmHg.</li>
	<li>At starting temperature, aim for a portal vein flow of 500-600 ml/min at 3-5 mmHg. After raising the temperature, monitor the portal venous flow, which will increase up to 1,100 ml/min at 4-6 mmHg. Avoid exceeding portal pressure above physiological values (around 8 mmHg) to protect sinusoidal fenestrations26. Avoid exceeding total flow above 2,000 ml/min in order to prevent damaging the organ. Set the outflow to -2 mmHg by lowering the main reservoir to prevent liver congestion by functional outflow obstruction.</li>
	<li>Add the dialysis component to the circuit in order to equilibrate the perfusion solution to predetermined values27. Set the dialysate flow to 500 ml/hr. Take special attention to adjust the dialysis outflow so that the perfusion solution is neither diluted nor concentrated. Within the first hour of perfusion the main reservoir must be watched carefully!</li>
	<li>Ensure homogeneous oxygenation of the tissue to recover and maintain organ function by using a main gas mixture component of O2 (95-98%) and CO2 (2-5%). Use variable gas during perfusion since the liver changes its metabolism and its pH demand during perfusion.</li>
	<li>Maintain a low pH during the start of perfusion to protect the organ using the paradox pH concept28 and avoid severe tissue damage that can result from fast connections to physiological pH under reoxygenation, since after storage in UW solution, the organ has an acidotic pH below 7. Adjust the partial pressure of CO2 continuously down to 25-30 mmHg so that the pH will reach a physiologic level within 1 hr.</li>
	<li>Add sodium- or potassium bicarbonate to the circuit to achieve a physiological concentration of standard bicarbonate in the perfusion solution. Inject it carefully under repetitive blood gas and electrolyte control.</li>
	<li>Monitor the perfusion by periodical venous and arterial blood gas and AST analyses. The venous PO2 remains above 175 mmHg during the perfusion. Monitor vascular flow and pressure and note a stable perfusion by a constant vascular resistance.</li>
	<li>Keep the perfusion system stable for up to 8 hr. At the end of the ex vivo perfusion period, cool down the perfusion system to 20 &#176;C and, after disconnecting the circuit&#8217;s tubing from the liver, flush the perfusion solution out the liver dually with ice cold UW solution. Store the liver once again placed on ice in a sterile organ bag.</li>
</ol>

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The authors have nothing to disclose.

In a pig model that mimics DCD liver transplantation, we demonstrated that subnormothermic liver perfusion with a cellular perfusion solution results in stable perfusion parameters, minimal hepatocyte injury, and active hepatic metabolism. Our subnormothermic perfusion set up has proven to recover a hepatocellular homeostasis and metabolism. Glycogen storage is restored and metabolites are discarded.
Ex vivo liver perfusion as preservation technique offers for the first time the opportunity to assess markers of graft function and injury during organ preservation and prior to transplantation. Beside the macroscopic evaluation of the graft perfusion homogeneity, flow values provide a good indicator of the graft&#8217;s viability and the extent of the ischemic injury it had suffered earlier29. Oxygen consumption and bile production are markers of metabolic function. Levels of hepatic enzymes like AST can be used to assess the degree and dynamics of hepatocellular injury30. This thorough graft assessment may allow a reliable discrimination between transplantable and non-transplantable marginal organs.
We chose a subnormothermic temperature of 33 &#176;C in our perfusion system because the temperature is sufficient to allow metabolism as well as ATP and glycogen synthesis. At the same time, it provides a decreased oxygen demand in comparison to normothermic perfusion settings which provides additional safety against ischemic injury. In general, perfusion temperatures above 30 &#176;C have shown to minimize cold ischemic injury and provide sufficient metabolic activity31.
Contrary to other groups, we did not use whole blood as perfusate, but a normo-osmotic albumin solution (Steen) with washed and filtered red blood cells. By excluding the plasma components as well as thrombocytes and leukocytes, the perfusion solution is designed to minimize pro-inflammatory signaling during the ex vivo perfusion.
In addition to the graft assessment, stable perfusion conditions over several hours allow graft treatment. Numerous molecules have shown to attenuate reperfusion injury under experimental conditions32. However, almost no treatment regime has made its way into clinical practice, yet. One reason seems to be the lack of opportunity to apply those treatments during cold storage. A metabolically active liver on an ex vivo perfusion system is optimal for applying any kind of treatment. In this regard, not only treatments to ameliorate reperfusion conditions like attenuation of Kupffer cell activity or scavenging of reactive oxygen species are conceivable but also treatments like gene therapy to condition the graft, e.g., against Hepatitis C recurrence. Other potential strategies could include reduction on steatosis during the ex vivo perfusion period33.
In summary, ex vivo liver perfusion is a novel strategy to minimize cold ischemic injury and to assess marginal liver grafts prior to liver transplantation. The ex vivo perfusion setting provides unique opportunity to repair and condition grafts prior to transplantation.

Liver transplantation is the only treatment option for patients with end stage liver disease or advanced hepatocellular carcinoma. For the last 25 years, the number of waiting list candidates has gradually increased and exceeded the number of available grafts. The number of heart beating donors has decreased over the last decade. At the same time, numbers of marginal grafts, such as donation after cardiac death (DCD), as well as old and fatty livers have increased1,2.
Marginal grafts are often declined for liver transplantation because of the higher chance of primary graft non- or delayed function. In DCD grafts, development of ischemic type biliary strictures (ITBS) is of special concern. With the conventional static cold preservation technique, ITBS occur in about 10-40% of DCD grafts. In the majority of patients, ITBS leads to re-transplantation or patient death. Especially prolonged warm and cold ischemic times are risk factors for ITBS3-7. Donor age, genetic predispositions (such as CCR5 delta 32), and the choice of preservation solution have also been discussed as additional risk factors7-10. Partial microthrombosis of the peribiliary vessels has been suggested as potential mechanism for ITBS after liver transplantation with DCD grafts11.
Prior to the clinical introduction of liver transplantation, ex vivo liver perfusions have been used to study hepatic metabolism and physiology12,13. After liver transplantation found its way into the clinical setting in the 1960s, innumerable attempts have been made to use ex vivo liver perfusion as a preservation method by mimicking physiological nutrition and oxygenation conditions. Its utility for preservation of marginal grafts has been investigated in the last decade, but it did not reach standard clinical care. We recently described a reduction in bile duct injury in DCD liver transplantation by ex vivo perfused preservation14. Different approaches regarding the perfusion solution were made. The selection ranges from cellular solutions like whole blood from the donor animal or packed red cells in combination with human plasma, to acellular approaches like machine University of Wisconsin solution, IGL solution, or Steen solution14-19.
The temperature ranges from 4-37 &#176;C20. The nomenclature in hypothermic, subnormothermic, and normothermic is very variable and inconsistent. All different techniques, solutions, and temperature settings aim at 1) stable perfusion conditions, 2) sufficient oxygenation, and 3) re-establishment of organ function. An enhanced preservation capacity as well as the ability of organ assessment and treatment during normothermic and subnormothermic perfusion faces higher technical complexity and costs compared to hypothermic perfusion20,21.
We have developed a subnormothermic ex vivo liver perfusion system over the last 4 years. The system can be used to 1) &#8220;recharge&#8221; the hepatic energy content, 2) to assess the graft&#8217;s quality, and to 3) repair marginal livers prior to transplantation. The following protocol contains all information for a stable hepatic perfusion.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>circuit</td>
			<td>Maquet (Hirrlingen, GER)</td>
			<td>custom made</td>
			<td>main reservoir (3L, 3/8&#34; outflow)</td>
		</tr>
		<tr>
			<td>-</td>
			<td></td>
			<td></td>
			<td>portal reservoir (1.5L, 1/4&#34;, outflow)</td>
		</tr>
		<tr>
			<td>-</td>
			<td></td>
			<td></td>
			<td>centrifugal pump</td>
		</tr>
		<tr>
			<td>-</td>
			<td></td>
			<td></td>
			<td>oxygenator</td>
		</tr>
		<tr>
			<td>-</td>
			<td></td>
			<td></td>
			<td>leukocyte filter</td>
		</tr>
		<tr>
			<td>tubing (1/4&#34; x 1/16&#34;)</td>
			<td>Raumedic (Helmbrechts, GER)</td>
			<td>MED7506</td>
			<td></td>
		</tr>
		<tr>
			<td>tubing (3/8&#34; x 3/32&#34;)</td>
			<td>Raumedic (Helmbrechts, GER)</td>
			<td>MED7536</td>
			<td></td>
		</tr>
		<tr>
			<td>tubing connectors</td>
			<td>Raumedic (Helmbrechts, GER)</td>
			<td></td>
			<td>various sizes</td>
		</tr>
		<tr>
			<td>dialysis filter, Optiflux F160NR</td>
			<td>Fresenius Medical Care (Waltham, MA)</td>
			<td>F160NR</td>
			<td></td>
		</tr>
		<tr>
			<td>STEEN solution</td>
			<td>XVIVO (G&#246;teborg, SWE)</td>
			<td>19004</td>
			<td>2L</td>
		</tr>
		<tr>
			<td>dialysis acid concentrate A</td>
			<td>Baxter (Mississauga, ON)</td>
			<td>D12188M</td>
			<td>45ml</td>
		</tr>
		<tr>
			<td>amino acid, Travasol 10%</td>
			<td>Baxter (Mississauga, ON)</td>
			<td>JB6760</td>
			<td>100ml</td>
		</tr>
		<tr>
			<td>Sodium Pyruvate</td>
			<td>Sigma-Aldrich (St. Louis, MO)</td>
			<td>P2256</td>
			<td>1.1g</td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>Sandoz Canada Inc (Toronto, ON)</td>
			<td>10750</td>
			<td>40000 iU</td>
		</tr>
		<tr>
			<td>Calcium Gluconate</td>
			<td>Pharmaceutical Partners of Canada (Richmond Hill, ON)</td>
			<td>C31110</td>
			<td>10mg</td>
		</tr>
		<tr>
			<td>fast acting Insulin</td>
			<td>various vendors</td>
			<td></td>
			<td>1000 iU</td>
		</tr>
		<tr>
			<td>Cefazoline</td>
			<td>various vendors</td>
			<td></td>
			<td>1g</td>
		</tr>
		<tr>
			<td>Metronidazole</td>
			<td>Baxter (Mississauga, ON)</td>
			<td>JB3401</td>
			<td>500mg</td>
		</tr>
	</tbody>
</table>

technique of subnormothermic ex vivo liver perfusion for the storage, assessment, and repair of marginal liver grafts

The success of liver transplantation has resulted in a dramatic organ shortage. In most transplant regions 20-30% of patients on the waiting list for liver transplantation die without receiving an organ transplant or are delisted for disease progression. One strategy to increase the donor pool is the utilization of marginal grafts, such as fatty livers, grafts from older donors, or donation after cardiac death (DCD). The current preservation technique of cold static storage is only poorly tolerated by marginal livers resulting in significant organ damage. In addition, cold static organ storage does not allow graft assessment or repair prior to transplantation.
These shortcomings of cold static preservation have triggered an interest in warm perfused organ preservation to reduce cold ischemic injury, assess liver grafts during preservation, and explore the opportunity to repair marginal livers prior to transplantation. The optimal pressure and flow conditions, perfusion temperature, composition of the perfusion solution and the need for an oxygen carrier has been controversial in the past.
In spite of promising results in several animal studies, the complexity and the costs have prevented a broader clinical application so far. Recently, with enhanced technology and a better understanding of liver physiology during ex vivo perfusion the outcome of warm liver perfusion has improved and consistently good results can be achieved.
This paper will provide information about liver retrieval, storage techniques, and isolated liver perfusion in pigs. We will illustrate a) the requirements to ensure sufficient oxygen supply to the organ, b) technical considerations about the perfusion machine and the perfusion solution, and c) biochemical aspects of isolated organs.

Below, we present the results of 5 perfusion experiments with DCD-grafts after 45 min warm- and 4 hr cold ischemia prior to the start of the subnormothermic ex vivo perfusion.
The main goal for an ex vivo liver perfusion is to ensure a sufficient oxygen supply to the organ. Ischemia causes vasoconstriction, thus increasing the perfusion resistance. Achieving constant vascular flows with stable pressures is a good indicator of adequate oxygenation. During an induction period of 1-2 hr the perfusion solution and the organ are warmed up to 33 &#176;C, which deceases the vascular resistance of the liver. Once the target temperature of 33 &#176;C is achieved, flow values level at a constant, nearly physiological range for the rest of the 6 hr perfusion time (Figures 3A-3D).
At the same time, the organ becomes metabolically active. Figure 4A shows the venous pO2, a marker of oxygen consumption. Within the initial 2 hr the venous pO2 declines to a constant plateau. At this metabolically active state, the liver starts producing bile (Figure 4B). The dialyzer provides a balanced electrolyte homeostasis (Figures 4C-4D). An initial hyperkalemia is quickly leveled out. Online AST measurement serves as monitoring of hepatocellular damage. Figure 5 displays only a shallow linear AST increase over the entire perfusion period. H&#38;E staining after 6 hr of perfusion reveals hepatocyte necrosis &#60;5 % with an intact lobular and sinusoidal structure (Figure 6). PAS staining at the same time point shows replenished cellular glycogen storage compared to exhausted storage in cold preserved DCD-grafts (Figure 7).
<img alt="Figure 3" src="/files/ftp_upload/51419/51419fig3highres.jpg" /> 
Figure 3. Perfusion flows and pressure (n = 5, error bars show standard deviation). (A,B) Hepatic artery (HA) flow and pressure: During the warming phase in the first 1-2 hr, the flow increases at stable pressures and is constant afterwards. Looking at the decreasing portal venous pressure (C), the increase of HA flow towards the end of the perfusion might be an autoregulatory reaction of the liver. (C,D) The portal venous (PV) flow increases corresponding to the HA flow during the first 2 hr of warming. The pressures remain relatively stable.
<img alt="Figure 4" src="/files/ftp_upload/51419/51419fig4highres.jpg" /> 
Figure 4. Monitoring parameters (n = 5, error bars show standard deviation). (A) The venous pO2 as a marker of oxygen demand and metabolic activity decreases within the initial phase of warming due to activated cellular metabolism; it remains stable afterwards. (B) Bile production as a marker of metabolic activity starts at temperatures around 30 &#176;C and, thus, between the first and second hour of perfusion. (C,D) The dialyzer assures electrolyte homeostasis; an initial hyperkalemia is quickly balanced.
<img alt="Figure 5" src="/files/ftp_upload/51419/51419fig5highres.jpg" /> 
Figure 5. AST (n = 5, error bars show standard deviation). AST is a sensitive marker of hepatocellular injury; the shallow increase suggests no significant injury during ex vivo perfusion.
<img alt="Figure 6" src="/files/ftp_upload/51419/51419fig6highres.jpg" /> 
Figure 6. H&#38;E staining (20X magnification). (A) Sham liver sample before warm ischemia, one representative liver lobule with intact architecture. (B) Liver sample after 45 min of warm ischemia, 4 hr of cold ischemia, and 6 hr of subnormothermic perfusion, the lobular architecture is intact without necrosis and only minimal cell swelling, the sinusoidal spaces are mildly dilated in comparison to the sham sample.

Watch this Scientific Journal Video about Technique of Subnormothermic Ex Vivo Liver Perfusion for the Storage, Assessment, and Repair of Marginal Liver Grafts at JoVE.com

Technique of Subnormothermic Ex Vivo Liver Perfusion for the Storage, Assessment, and Repair of Marginal Liver Grafts

Marginal grafts, such as fatty livers, grafts from older donors, or livers retrieved after cardiac death (DCD) tolerate conventional, cold static storage only poorly. We developed a novel model of subnormothermic ex vivo liver perfusion for preservation, assessment, and repair of marginal liver grafts prior to transplantation.

Technique of Subnormothermic Ex Vivo Liver Perfusion for the Storage, Assessment, and Repair of Marginal Liver Grafts

The success of liver transplantation has resulted in a dramatic organ shortage. In most transplant regions 20-30% of patients on the waiting list for ...

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14.7K Views.  Toronto General Hospital.  Предельные трансплантаты, такие как жирные печени, трансплантатов из более старых доноров или печени, получаемых после сердечной смерти (DCD) переносят обычную, холодный статического хранения только слабо. Мы разработали новую модель subnormothermic экс естествен?...

Video: Technique of Subnormothermic Ex Vivo Liver Perfusion for the Storage, Assessment, and Repair of Marginal Liver Grafts

Steps for the Autologous Ex vivo Perfused Porcine Liver-kidney Experiment

The use of ex vivo perfused models can mimic the physiological conditions of the liver for short periods, but to maintain normal homeostasis for an extended perfusion period is challenging. We have added the kidney to our previous ex vivo perfused liver experiment model to reproduce a more accurate physiological state for prolonged experiments without using live animals. Five intact livers and kidneys were retrieved post-mortem from sacrificed pigs on different days and perfused for a minimum of 6 hr. Hourly arterial blood gases were obtained to analyze pH, lactate, glucose and renal parameters. The primary endpoint was to investigate the effect of adding one kidney to the model on the acid base balance, glucose, and electrolyte levels. The result of this liver-kidney experiment was compared to the results of five previous liver only perfusion models. In summary, with the addition of one kidney to the ex vivo liver circuit, hyperglycemia and metabolic acidosis were improved. In addition this model reproduces the physiological and metabolic responses of the liver sufficiently accurately to obviate the need for the use of live animals. The ex vivo liver-kidney perfusion model can be used as an alternative method in organ specific studies. It provides a disconnection from numerous systemic influences and allows specific and accurate adjustments of arterial and venous pressures and flow.

Steps for the Autologous Ex vivo Perfused Porcine Liver-kidney Experiment

Study of the animal organ physiology can be achieved by performing an experimental ex vivo perfusion system. The addition of a porcine kidney, as a homeostatic organ to our previously developed ex vivo liver perfusion model can be a principal step to achieve a better physiological environment.

The use of ex vivo perfused models can mimic the physiological conditions of the liver for short periods, but to maintain normal homeostasis for an ...

Isolated Hepatic Perfusion as a Treatment for Liver Metastases of Uveal Melanoma

Isolated hepatic perfusion (IHP) is a procedure where the liver is surgically isolated and perfused with a high concentration of the chemotherapeutic agent melphalan. Briefly, the procedure starts with the setup of a percutaneous veno-venous bypass from the femoral vein to the external jugular vein. Via a laparotomy, catheters are then inserted into the proper hepatic artery and the caval vein. The portal vein and the caval vein, both supra- and infrahepatically, are then clamped. The arterial and venous catheters are connected to a heart lung machine and the liver is perfused with melphalan (1 mg/kg body weight) for 60 min. This way it is possible to locally perfuse the liver with a high dose of a chemotherapeutic agent, without leakage to the systemic circulation.
In previous studies including patients with isolated liver metastases of uveal melanoma, an overall response rate of 33-100% and a median survival between 9 and 13 months, have been reported. The aim of this protocol is to give a clear description of how to perform the procedure and to discuss IHP as a treatment option for liver metastases of uveal melanoma.

Here, we present a protocol how to perform an isolated liver perfusion (IHP) with melphalan and we also discuss IHP as a treatment option for liver metastases of uveal melanoma.

Isolated hepatic perfusion (IHP) is a procedure where the liver is surgically isolated and perfused with a high concentration of the chemotherapeutic ...

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 ...

Functional Human Liver Preservation and Recovery by Means of Subnormothermic Machine Perfusion

There is currently a severe shortage of liver grafts available for transplantation. Novel organ preservation techniques are needed to expand the pool of donor livers. Machine perfusion of donor liver grafts is an alternative to traditional cold storage of livers and holds much promise as a modality to expand the donor organ pool. We have recently described the potential benefit of subnormothermic machine perfusion of human livers. Machine perfused livers showed improving function and restoration of tissue ATP levels. Additionally, machine perfusion of liver grafts at subnormothermic temperatures allows for objective assessment of the functionality and suitability of a liver for transplantation. In these ways a great many livers that were previously discarded due to their suboptimal quality can be rescued via the restorative effects of machine perfusion and utilized for transplantation. Here we describe this technique of subnormothermic machine perfusion in detail. Human liver grafts allocated for research are perfused via the hepatic artery and portal vein with an acellular oxygenated perfusate at 21 &#176;C.

We describe a method of ex vivo machine perfusion of human liver grafts at subnormothermic temperature (21 &#176;C).

There is currently a severe shortage of liver grafts available for transplantation.  Novel organ preservation techniques are needed to expand the pool of ...

Normothermic Ex Vivo Kidney Perfusion for the Preservation of Kidney Grafts prior to Transplantation

Kidney transplantation has become a well-established treatment option for patients with end-stage renal failure. The persisting organ shortage remains a serious problem. Therefore, the acceptance criteria for organ donors have been extended leading to the usage of marginal kidney grafts. These marginal organs tolerate cold storage poorly resulting in increased preservation injury and higher rates of delayed graft function. To overcome the limitations of cold storage, extensive research is focused on alternative normothermic preservation methods.
Ex vivo normothermic organ perfusion is an innovative preservation technique. The first experimental and clinical trials for ex vivo lung, liver, and kidney perfusions demonstrated favorable outcomes.
In addition to the reduction of cold ischemic injury, the method of normothermic kidney storage offers the opportunity for organ assessment and repair. This manuscript provides information about kidney retrieval, organ preservation techniques, and isolated ex vivo normothermic kidney perfusion (NEVKP) in a porcine model. Surgical techniques, set up for the perfusion solution and the circuit, potential assessment options, and representative results are demonstrated.

Normothermic Ex Vivo Kidney Perfusion for the Preservation of Kidney Grafts prior to Transplantation

The severe organ shortage has resulted in increased use of marginal kidney grafts for transplantation. This has triggered interest in alternative storage methods, since marginal grafts especially tolerate cold storage poorly. The technique of normothermic ex vivo kidney perfusion (NEVKP) represents a novel preservation method for kidney grafts prior to &#160;transplantation.

Kidney transplantation has become a well-established treatment option for patients with end-stage renal failure. The persisting organ shortage remains a ...

Method of Direct Segmental Intra-hepatic Delivery Using a Rat Liver Hilar Clamp Model

Major hepatic surgery with inflow occlusion, and liver transplantation, necessitate a period of warm ischemia, and a period of reperfusion leading to ischemia/reperfusion (I/R) injury with myriad negative consequences. Potential I/R injury in marginal organs destined for liver transplantation contributes to the current donor shortage secondary to a decreased organ utilization rate. A significant need exists to explore hepatic I/R injury in order to mediate its impact on graft function in transplantation. Rat liver hilar clamp models are used to investigate the impact of different molecules on hepatic I/R injury. Depending on the model, these molecules have been delivered using inhalation, epidural infusion, intraperitoneal injection, intravenous administration or injection into the peripheral superior mesenteric vein. A rat liver hilar clamp model has been developed for use in studying the impact of pharmacologic molecules in ameliorating I/R injury. The described model for rat liver hilar clamp includes direct cannulation of the portal supply to the ischemic hepatic segment via a side branch of the portal vein, allowing for direct segmental hepatic delivery. Our approach is to induce ischemia in the left lateral and median lobes for 60 min, during which time the substance under study is infused. In this case, pegylated-superoxide dismutase (PEG-SOD), a free radical scavenger, is infused directly into the ischemic segment. This series of experiments demonstrates that infusion of PEG-SOD is protective against hepatic I/R injury. Advantages of this approach include direct injection of the molecule into the ischemic segment with consequent decrease in volume of distribution and reduction in systemic side effects.

A unique rat liver hilar clamp model was developed for studying the impact of pharmacologic molecules in ameliorating ischemia-reperfusion injury. This model includes direct cannulation of the portal supply to the ischemic liver segment via a branch of the portal vein, allowing for direct hepatic delivery.

Major hepatic surgery with inflow occlusion, and liver transplantation, necessitate a period of warm ischemia, and a period of reperfusion leading to ...

Assessment of Plasma Coagulation on Liver Tissue in a Large Animal Model In Vivo

Plasma coagulation as a form of electrocautery is used in liver surgery for decades to seal the large liver cut surface after major hepatectomy to prevent hemorrhages at a later stage. The exact effects of plasma coagulation on liver tissue are only poorly examined. In our porcine model, the coagulation effects can be examined close to the clinical application. A combined laser Doppler flowmeter and spectrophotometer documents microcirculation changes during coagulation at 8 mm tissue depth noninvasively, providing quantifiable information about hemostasis beyond the subjective clinical impression. The temperature at coagulation site is assessed with an infrared thermometer prior and post coagulation and with a thermographic camera during coagulation, a measurement of the gas beam temperature is not possible due to the upper threshold of the devices. The depth of coagulation is measured microscopically on hematoxylin/eosin stained sections after calibration with an object micrometer and gives an exact information about the power setting-coagulation depth-relation. The sealing effect is examined on the bile ducts as it is not possible for a plasma coagulator to seal larger blood vessels. Burst pressure experiments are carried out on explanted organs to rule out blood pressure related effects.

Assessment of Plasma Coagulation on Liver Tissue in a Large Animal Model In Vivo

Here we present a protocol to experimentally assess plasma coagulation in liver tissue in vivo. In a porcine model, microcirculation is examined by laser Doppler, coagulation depth is measured histologically, temperature at coagulation site by infrared thermometer and thermographic camera, and duct sealing effect is documented by burst pressure experiments.

Plasma coagulation as a form of electrocautery is used in liver surgery for decades to seal the large liver cut surface after major hepatectomy to prevent ...

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, normothermic ex vivo liver perfusion (NEVLP) has been introduced as a method to evaluate and modify organ function. NEVLP has many advantages in comparison to hypothermic and subnormothermic perfusion including reduced preservation injury, restoration of normal organ function under physiologic conditions, assessment of organ performance, and as a platform for organ repair, remodeling, and modification. Both murine and porcine NEVLP models have been described. We demonstrate a rat model of NEVLP and use this model to show one of its important applications &#8212; the use of a therapeutic molecule added to liver perfusate. Catalase is an endogenous reactive oxygen species (ROS) scavenger and has been demonstrated to decrease ischemia-reperfusion in the eye, brain, and lung. Pegylation has been shown to target catalase to the endothelium. Here, we added pegylated-catalase (PEG-CAT) to the base perfusate and demonstrated its ability to mitigate liver preservation injury. An advantage of our rodent NEVLP model is that it is inexpensive in comparison to larger animal models. A limitation of this study is that it does not currently include post-perfusion liver transplantation. Therefore, prediction of the function of the organ post-transplantation cannot be made with certainty. However, the rat liver transplant model is well established and certainly could be used in conjunction with this model. In conclusion, we have demonstrated an inexpensive, simple, easily replicable NEVLP model using rats. Applications of this model can include testing novel perfusates and perfusate additives, testing software designed for organ evaluation, and experiments designed to repair organs.

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant liver donor shortage, and criteria for liver donors have been expanded. Normothermic ex vivo liver perfusion (NEVLP) has been developed to evaluate and modify organ function. This study demonstrates a rat model of NEVLP and tests the ability of pegylated-catalase, to mitigate liver preservation injury.

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, ...

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 ...

Normothermic Ex Vivo Pancreas Perfusion for the Preservation of Pancreas Allografts before Transplantation

Pancreas transplantation (PTx) is a curative treatment for people who live with the burden of a diagnosis of diabetes mellitus (DM). However, due to organ shortages and increasing numbers of patients being listed for PTx, new strategies are needed to increase the number of available grafts for transplantation. 
Static cold storage (SCS) is considered the gold standard for standard criteria organs. However, standard criteria donors (SCD) are becoming scarce and new strategies that can increase the rate of organ acceptance from extended criteria donors (ECD) are urgently needed. 
Normothermic ex vivo perfusion (NEVP) is one of the strategies that has become increasingly popular over the past couple of decades. This preservation method has already been used successfully in other organs (liver, kidneys, and lungs) but has been minimally explored in pancreas transplantation. The few papers that describe the method for pancreas show little success, edema being one of the major issues. The following manuscript describes the successful NEVP method and setup developed by our group to perfuse swine pancreas.

Normothermic Ex Vivo Pancreas Perfusion for the Preservation of Pancreas Allografts before Transplantation

Normothermic ex vivo machine perfusion (NEVP) has scarcely been explored for the preservation of pancreas allografts. We present an innovative preservation technique for pancreas allografts before transplantation.

Pancreas transplantation (PTx) is a curative treatment for people who live with the burden of a diagnosis of diabetes mellitus (DM). However, due to organ ...