Throughout the writing of this paper, I have received a great deal of support and assistance. I would particularly like to acknowledge my teammate XinPei Chen for his wonderful collaboration and patient support during my operation.

Animal experiments were performed according to the current German regulations and guidelines for animal welfare and the ARRIVE guidelines for Reporting Animal Research. The animal experiment protocol was approved by the Th&#252;ringer Landesamt f&#252;r Verbraucherschutz, Thuringia, Germany (Approval-Number: UKJ - 17 - 106).
NOTE: Male C57BL/6J mice weighing 34 &#177; 4 g (mean &#177; standard error of the mean [SEM]) were used as liver donors. They were maintained under controlled environmental conditions (50% humidity and 18 - 23 &#176;C) with free access to standard mouse chow and water. Throughout the surgical procedure, a respiratory rate exceeding 60 breaths/min was maintained, and body temperature was kept above 34 &#176;C.
1. Preparation
<ol>
	<li>Setting up the operation table
	<ol>
		<li>Autoclave all surgical instruments and consumables for sterilization purposes.</li>
		<li>Turn on all equipment, including the warming board and electrocoagulation.</li>
		<li>Place one 50 mL syringe with 25 mL heparinized (2,500 U/L) saline in a warm incubator (37 &#176;C).</li>
		<li>Place the surgical instruments, the 6 - 0 silk suture, sterile small cotton applicator, veterinary saline (500 mL), and non-woven gauze sponges (10 cm x 10 cm) on the operation table appropriately.</li>
		<li>Place a 26 G needle on the operation table to create a small hole in the lid of the 0.5 mL microcentrifuge tube to receive the biliary tube for bile collection.</li>
		<li>Place the cannula (1 Fr polyurethane cannula or UT - 03 polyethylene cannula) and a sterilized 0.5 mL microcentrifuge tube for bile collection on the operation table.</li>
	</ol>
	</li>
	<li>Self-made portal vein cannula
	<ol>
		<li>Hold the 2 Fr cannula with forceps and puncture the wall with a 30 G needle at a 1 cm distance from the end of the cannula. Push the needle through the cannula until the tip of the needle becomes visible.</li>
		<li>Trim the tip of the cannula resulting in a sharp triangle.</li>
	</ol>
	</li>
	<li>Preparation of heparinized saline
	<ol>
		<li>Prepare 25 mL of heparinized saline with a final concentration of 2,500 IU/mL.</li>
		<li>Remove all air bubbles and place the syringe in the 40 &#176;C incubator.</li>
	</ol>
	</li>
	<li>Demonstration of the perfusion system
	<ol>
		<li>See Figure 1 for the main components of the machine perfusion system.</li>
	</ol>
	</li>
	<li>Set up of the organ chamber
	<ol>
		<li>See Figure 2 for the layout of the organ chamber.</li>
	</ol>
	</li>
	<li>Set up of the perfusion system
	<ol>
		<li>Turn on the lab chart program for pressure monitoring.</li>
		<li>Connect the pressure calibrator and pressure sensor at the organ chamber level.</li>
		<li>Adjust the pressure calibrator to read 0 mmHg and check the corresponding value on the pressure control software.</li>
		<li>Adjust the pressure calibrator to read 20 mmHg and again check the corresponding value on the pressure control software.</li>
		<li>Turn on the water bath, and prewarm the organ chamber to 40 &#176;C.</li>
		<li>Flush the entire plumbing system twice with distilled deionized water for 30 min each, ensuring complete removal of the sanitizing solution.</li>
		<li>Initiate the circulation of the disinfection solution throughout the entire system for a duration of 20 min to ensure thorough disinfection.</li>
		<li>Turn on the gas mixture (95% oxygen (O2) and 5% carbon dioxide (CO2).</li>
	</ol>
	</li>
	<li>Perfusate filling
	<ol>
		<li>Supplement 250 mL of Williams&#39; E medium with 50 mL of fetal bovine serum, 3 mL of penicillin/ streptomycin (1 mg/mL), 0.17 mL of insulin (100 IE/mL), 0.34 mL of heparin (5000 U/mL), and 0.07 mL of hydrocortisone (100 mg/2 mL) to prepare the complete Williams&#39; E medium.</li>
		<li>Add equal volumes (150 mL) of perfusate to the reservoir and the organ chamber to prime the system. 
		NOTE: Special attention must be paid to maintain sterility during the filling process. The perfusate is constantly pumped through these two key components of the closed recirculating machine perfusion.</li>
		<li>Turn on the peristaltic pump at medium speed (15 mL/min) to prime the perfusion system with the oxygenated medium.</li>
	</ol>
	</li>
</ol>
2. Liver explantation
<ol>
	<li>Pre-surgery preparation
	<ol>
		<li>Weigh the animal. Prepare the analgesic buprenorphine (0.3 mg/mL) (0.05 mg/kg body weight).</li>
		<li>Connect the induction chamber with the wall socket. Turn oxygen to 0.5 L/min. Turn isoflurane to 3%.</li>
		<li>Place the animal in the chamber until deep anesthesia (righting reflex positive) is reached.</li>
		<li>Use a micro syringe to apply body weight-adapted dose of analgesia subcutaneously.</li>
		<li>Use an electric shaver to trim the fur on the abdominal skin.</li>
		<li>Transfer the mouse to the operation table and turn on the isoflurane vaporizer to 2.5% to maintain anesthesia. Confirm the depth of anesthesia by testing the interdigital toe reflex.</li>
	</ol>
	</li>
	<li>Preparation of the mouse abdomen
	<ol>
		<li>Place the mouse in a supine position.</li>
		<li>Test interdigital reflex to double confirm the appropriate depth of anesthesia. Fix all four limbs with tape.</li>
		<li>Disinfect both sides of the abdomen to the mid-axillary line using three consecutive rounds of iodine-alcohol. Use non-woven sterilized gauze to cover the area around the surgical field.</li>
		<li>Make a 3 cm transverse incision 1 cm below the xiphoid in the abdominal area of the mouse using Metzenbaum baby scissors and surgical forceps.</li>
		<li>Extend the skin incision bilaterally to the midaxillary line on both sides.</li>
		<li>Carefully make a 2 cm longitudinal incision along the linea alba using spring scissors.</li>
		<li>Cut through the abdominal muscle layer with electrocoagulation and Vannas spring scissors.</li>
		<li>Carefully place a piece of wet gauze to protect the liver from electrocoagulation.</li>
		<li>Use a 6 - 0 silk suture with the round needle to retract the xiphoid process for better exposure of the coronary ligament.</li>
		<li>Use two rib retractors to fully expose the abdominal cavity of the mouse.</li>
		<li>Carefully move the small intestine out of the abdominal cavity with a wet cotton swab to fully expose the hilum.</li>
	</ol>
	</li>
	<li>Common bile duct preparation
	<ol>
		<li>Transect the falciform, phrenic, and gastrohepatic ligaments with sharp scissors.</li>
		<li>Carefully free the common bile duct using fine curved forceps without teeth. 
		&#8203;NOTE: The common bile duct is very easily damaged and breaks. Once it breaks, it cannot be cannulated. Due to the direction of the anatomical position, curved forceps are better to be used.</li>
		<li>Place two 6 - 0 silk suture loops over the common bile duct in preparation for the next step.</li>
	</ol>
	</li>
	<li>Common bile duct cannulation
	<ol>
		<li>Carefully puncture the bile duct with a 30 G needle. Use pointed curved forceps to enlarge the small hole to fit the bile duct cannulation.</li>
		<li>Use vessel cannulation forceps to grasp the bile duct cannula and push it into the bile duct.</li>
		<li>Double secure the cannula with the preset 6 - 0 suture loops. 
		NOTE: During the cannulation, resistance by bile is felt. If the force is not well controlled, the cannula will be pushed out of the biliary tract by the pressure of bile outflow. Carefully adjust the depth of the cannula. If it is too deep, it may damage the bile duct, and if it is not deep enough, it may slip out.</li>
		<li>Observe the bile flow in the cannula after successful cannulation.</li>
	</ol>
	</li>
	<li>Portal vein preparation
	<ol>
		<li>Clamp the portal vein with flat forceps and carefully free the connective tissue with curved forceps. Do not pull hard to avoid causing tearing of the portal vein. Once the portal vein is damaged, it is difficult to re-cannulate the portal vein.</li>
		<li>Dissect the PV just superior to the bifurcation and place the first suture loop using 6 - 0 silk suture on PV close to the confluence for later use.</li>
		<li>Place the second suture loop for later fixation of the PV as close to the hepatic hilum as possible.</li>
	</ol>
	</li>
	<li>Portal vein cannulation
	<ol>
		<li>Use an arterial clip to close the distal portal vein.</li>
		<li>Very carefully, puncture the portal vein with one of the above portal vein cannulas. Blood flow can be clearly observed within the cannula after a successful puncture.</li>
		<li>Secure the PV cannula with the pre-placed 6 - 0 suture loop.</li>
	</ol>
	</li>
	<li>Liver flushing
	<ol>
		<li>Increase isoflurane to 5% and euthanize the mouse with an overdose of isoflurane inhalation.</li>
		<li>Take prewarmed heparin saline solution from the incubator. Remove all air bubbles formed inside the heparinized saline.</li>
		<li>Fix the syringe with prewarmed heparinized saline into the syringe pump.</li>
		<li>Connect the extension tube of the syringe pump to the cannula of the portal vein, adjust the speed to 2 mL/min, and start the liver flushing.</li>
		<li>Observe the color of the liver at the end of the flushing procedure. Excise the liver once the color turns to a homogenous yellow.</li>
		<li>Transect the diaphragm, suprahepatic inferior vena cava, infra hepatic vena cava, hepatic artery, distal portal vein, and any remaining connective tissue.</li>
		<li>Place the liver into the Petri dish.</li>
	</ol>
	</li>
</ol>
3. Liver and chamber connection
<ol>
	<li>Liver transfer
	<ol>
		<li>Carefully transfer the liver into the organ chamber using a Petri dish.</li>
		<li>Keep a small amount of saline in the Petri dish to prevent the liver from drying out. 
		NOTE: Portal vein and bile duct can easily be twisted during this procedure, which may affect liver perfusion and bile collection.</li>
	</ol>
	</li>
	<li>Portal vein cannula connection
	<ol>
		<li>Slowly infuse normal saline into the portal vein cannula with a syringe to evacuate the air bubbles in the cannula.</li>
		<li>Connect the portal vein cannula into the perfusate outflow tube in the organ chamber.</li>
	</ol>
	</li>
	<li>Bile duct cannula connection
	<ol>
		<li>Guide the mouse bile duct cannula through the valve of a rubber cap which is connected to the organ chamber.</li>
		<li>Insert the bile duct cannula into a pre-prepared 0.5 mL microtube with a small hole in the lid.</li>
		<li>Place the microtube on clay outside of the organ chamber.</li>
	</ol>
	</li>
</ol>
4. Adjust the flow rate according to PV pressure
<ol>
	<li>Turn on the peristaltic pump from 1 mL/min.</li>
	<li>Check the portal vein pressure reading to adjust the flow rate.</li>
	<li>Maintain the portal vein pressure in the physiological range between 7 - 10 mmHg by adjusting the flow rate. 
	NOTE: Nominal flow rate can vary slightly depending on the usage and positioning of tubes.</li>
</ol>
5. Sample collection
<ol>
	<li>Obtain inlet perfusate samples from the portal vein inflow tube and outlet perfusate samples from the organ chamber in 3h intervals.</li>
	<li>Collect samples from all liver lobes for histological analysis at the end of the perfusion period of 12 h.</li>
</ol>

There are no financial conflicts of interest to disclose.

Critical steps in the protocol 
The two crucial steps in liver explantation are the cannulation of the portal vein (PV) and the subsequent cannulation of the bile duct (BD). These steps are of paramount importance in ensuring successful organ retrieval and subsequent perfusion or transplantation procedures.
Challenges and solutions 
PV cannulation presents three challenges: injury of the vessel wall, displacement of the catheter, and practicability o...

Liver transplantation represents the gold standard treatment for individuals with end-stage liver disease. Regrettably, the demand for donor organs surpasses the available supply, leading to a significant shortage. In 2021, approximately 24,936 patients were on the waiting list for a liver graft, while only 9,234 transplants were successfully performed1. The significant disparity between the supply and demand of liver grafts highlights the pressing necessity to investigate alternative strategies to broaden the donor pool and enhance the accessibility of liver grafts. One way of expanding the donor pool is to use marginal donors2. Marginal donors include those with advanced age, moderate or severe steatosis. Although the transplantation of marginal organs may yield favorable outcomes, the overall results remain suboptimal. As a result, the development of therapeutic strategies aimed at enhancing the function of marginal donors is currently underway3,4.
One of the strategies is to use machine perfusion, especially normothermic oxygenated machine perfusion, to improve the function of these marginal organs5. However, there is still a limited understanding of the molecular mechanisms that underlie the beneficial effects of normothermic oxygenated machine perfusion (NEVLP). Mice, with their abundant availability of genetically modified strains, serve as valuable models for investigating molecular pathways. For instance, the significance of autophagy pathways in mitigating hepatic ischemia-reperfusion injury has been increasingly recognized6,7. One important molecular pathway in the hepatic ischemia-reperfusion injury is the miR-20b-5p/ATG7 pathway8. Currently, there are a number of ATG knockout and conditional knock-out mouse strains available but no corresponding rat strains9.
Based on this background, the aim was to generate a miniaturized NEVLP platform for mouse liver grafts. This platform would facilitate the exploration and evaluation of potential genetically modified strategies aimed at improving the functionality of the donor&#39;s liver. Additionally, it was essential for the system to be suitable for long-term perfusion, enabling the ex vivo treatment of the liver, commonly referred to as &#34;organ repair.&#34;
Considering the limited availability of relevant in vitro data on mouse liver perfusion, the literature review focused on studies conducted in rats. A systematic search of literature spanning from 2010 to 2022 was performed using keywords such as &#34;normothermic liver perfusion,&#34; &#34;ex vivo or in vitro,&#34; and &#34;rats&#34;. This search aimed to identify optimal conditions in rodents, allowing us to determine the most appropriate approach.
The perfusion system consists of a sealed water-jacketed glass buffer reservoir, a peristaltic roller pump, an oxygenator, a bubble trap, a heat exchanger, an organ chamber, and a closed cycling tubing system (Figure 1). The system ensures precise maintenance of a constant perfusion temperature of 37 &#176;C using a dedicated thermo-static machine. The peristaltic roller pump drives the flow of the perfusate throughout the circuit. The perfusion circuit initiates at the insulated water-jacketed reservoir. Subsequently, the perfusate is directed through the oxygenator, which receives a gas mixture of 95% oxygen and 5% carbon dioxide from a dedicated gas bottle. Following oxygenation, the perfusate passes through the bubble trap, wherein any entrapped bubbles are redirected back to the reservoir by the peristaltic pump. The remaining perfusate flows through the heat exchanger and enters the organ chamber, from where it returns to the reservoir.
Here, we report our experiences establishing a NEVLP for mouse livers and share the promising results of a pilot experiment performed using the oxygenated medium without oxygen carriers.

<table><tbody><tr><td>0.5 ml Micro Tube PP</td><td>Sarstedt</td><td>72699</td><td></td></tr><tr><td>1 Fr Rubber Cannula</td><td>Vygon</td><td>Sample Cannula</td><td></td></tr><tr><td>10 &amp;micro;L Micro Syringe</td><td>Hamilton</td><td>701N</td><td></td></tr><tr><td>2 Fr Rubber Cannula</td><td>Vygon</td><td>Sample Cannula</td><td></td></tr><tr><td>24 G Butterfly Cannula</td><td>Terumo</td><td>SR+OF2419</td><td></td></tr><tr><td>26 G Butterfly Cannula</td><td>Terumo</td><td>SR+DU2619WX</td><td></td></tr><tr><td>30 G Hypodermic Needle</td><td>Sterican</td><td>100246</td><td></td></tr><tr><td>50 ml Syringe Pump</td><td>Braun</td><td>110356</td><td></td></tr><tr><td>6-0 Perma-Hand Seide</td><td>Ethicon</td><td>639H</td><td></td></tr><tr><td>Arterial Clip</td><td>Braun</td><td>BH014R</td><td></td></tr><tr><td>Autoclavable Moist Chamber</td><td>Hugo Sachs Elektronik</td><td>73-4733</td><td></td></tr><tr><td>Big Cotton Applicator&amp;nbsp;</td><td>NOBA Verbandmittel Danz GmbH</td><td>974018</td><td></td></tr><tr><td>Bubble Trap</td><td>Hugo-Sachs-Elektronik</td><td>V83163</td><td></td></tr><tr><td>Buprenovet (0.3 mg / ml)</td><td>Elanco</td><td>/</td><td></td></tr><tr><td>CIDEX OPA solution (2 L)</td><td>Cilag GmbH</td><td>20391</td><td></td></tr><tr><td>Electrosurgical Unit for Monopolar Cutting VIO&amp;reg; 50 C</td><td>ERBE</td><td>/</td><td></td></tr><tr><td>Fetal Bovine Serum(500 ml)&amp;nbsp;</td><td>Sigma-Aldrich</td><td>F7524-500ML</td><td></td></tr><tr><td>Gas Mixture (95 % oxygen &amp;amp; 5 % carbon dioxide)</td><td>House Supply</td><td>/</td><td></td></tr><tr><td>Heating Circulating Baths</td><td>Harvard-Apparatus</td><td>75-0310</td><td></td></tr><tr><td>Heparin 5000 (I.E. /5 ml)</td><td>Braun</td><td>1708.00.00</td><td></td></tr><tr><td>Hydrocortisone (100 mg / 2 ml)</td><td>Pfizer</td><td>15427276</td><td></td></tr><tr><td>Insulin(100 IE / ml)</td><td>Sigma</td><td>I0516-5ML</td><td></td></tr><tr><td>Iris Scissors&amp;nbsp;</td><td>Fine Science Instruments</td><td>15000-03</td><td></td></tr><tr><td>Isofluran (250 ml)</td><td>Cp-Pharma</td><td>1214</td><td></td></tr><tr><td>Membrane Oxygenator</td><td>Hugo Sachs Elektronik</td><td>T18728</td><td></td></tr><tr><td>Microsurgery Microscope&amp;nbsp;</td><td>Leica</td><td>M60</td><td></td></tr><tr><td>Mouse Retractor Set&amp;nbsp;</td><td>Carfil Quality</td><td>180000056</td><td></td></tr><tr><td>NanoZoomer 2.0 HT</td><td>Hamamatsu</td><td>/</td><td></td></tr><tr><td>Non-Woven Sponges&amp;nbsp;</td><td>Kompressen</td><td>866110</td><td></td></tr><tr><td>Penicillin Streptomycin (1 mg / ml)&amp;nbsp;</td><td>C.C.Pro</td><td>Z-13-M</td><td></td></tr><tr><td>Perfusion Extension Tube (30 cm)</td><td>Braun</td><td>4256000</td><td></td></tr><tr><td>Peristaltic Pump</td><td>Harvard-Apparatus</td><td>P-70</td><td></td></tr><tr><td>Petri Dishc 100x15 mm</td><td>VWR&amp;reg;</td><td>391-0578</td><td></td></tr><tr><td>Povidon-Jod (Vet-Sep Spray)</td><td>Livisto</td><td>799-416</td><td></td></tr><tr><td>Pressure Transducer Simulator</td><td>UTAH Medical Products</td><td>650-950</td><td></td></tr><tr><td>Reusable Blood Pressure Transducers</td><td>AD Instruments</td><td>MLT-0380/D</td><td></td></tr><tr><td>S &amp;amp; T Vessel Cannulation Forceps</td><td>Fine Science Instruments</td><td>00608-11</td><td></td></tr><tr><td>Small Cotton Applicator</td><td>NOBA Verbandmittel Danz GmbH</td><td>974116</td><td></td></tr><tr><td>Straight Forceps 10 cm&amp;nbsp;</td><td>Fine Science Instruments</td><td>00632-11</td><td></td></tr><tr><td>Suture Tying Forceps</td><td>Fine Science Instruments</td><td>11063-07</td><td></td></tr><tr><td>Syringe 50ml Original Perfusor</td><td>Braun</td><td>8728810F-06</td><td></td></tr><tr><td>UT - 03 Cannula</td><td>Unique Medical, Japan</td><td>/</td><td></td></tr><tr><td>Vannas Spring Scissors</td><td>Fine Science Instruments</td><td>15018-10</td><td></td></tr><tr><td>Veterinary Saline (500 ml)</td><td>WDT</td><td>18X1807</td><td></td></tr><tr><td>Water Jacketed Reservoir&amp;nbsp; 2 L</td><td>Harvard-Apparatus</td><td>73-3441</td><td></td></tr><tr><td>William&#039;s E Medium (500 ML)</td><td>Thermofischer Scientific</td><td>A1217601</td><td></td></tr></tbody></table>

normothermic ex vivo liver machine perfusion in mouse

This protocol presents an optimized erythrocytes-free NEVLP system using mouse livers. Ex vivo preservation of mouse livers was achieved by employing modified cannulas and techniques adapted from conventional commercial ex vivo perfusion equipment. The system was utilized to evaluate the preservation outcomes following 12 h of perfusion. C57BL/6J mice served as liver donors, and the livers were explanted by cannulating the portal vein (PV) and bile duct (BD), and subsequently flushing the organ with warm (37 &#176;C) heparinized saline. Then, the explanted livers were transferred to the perfusion chamber and subjected to normothermic oxygenated machine perfusion (NEVLP). Inlet and outlet perfusate samples were collected at 3 h intervals for perfusate analysis. Upon completion of the perfusion, liver samples were obtained for histological analysis, with morphological integrity assessed using modified Suzuki-Score through Hematoxylin-Eosin (HE) staining. The optimization experiments yielded the following findings: (1) mice weighing over 30 g were deemed more suitable for the experiment due to the larger size of their bile duct (BD). (2) a 2 Fr (outer diameter = 0.66 mm) polyurethane cannula was better suited for cannulating the portal vein (PV) when compared to a polypropylene cannula. This was attributed to the polyurethane material&#39;s enhanced grip, resulting in reduced catheter slippage during the transfer from the body to the organ chamber. (3) for cannulation of the bile duct (BD), a 1 Fr (outer diameter = 0.33 mm) polyurethane cannula was found to be more effective compared to the polypropylene UT - 03 (outer diameter = 0.30 mm) cannula. With this optimized protocol, mouse livers were successfully preserved for a duration of 12 h without significant impact on the histological structure. Hematoxylin-Eosin (HE) staining revealed a well-preserved morphological architecture of the liver, characterized by predominantly viable hepatocytes with clearly visible nuclei and mild dilation of hepatic sinusoids.

Establishment of surgical procedure 
A total of 17 animals were utilized for this experiment: 14 mice were employed for optimizing the organ procurement process, including cannulation of the portal vein (PV) and bile duct (BD), while 3 mice were used to validate the procedure (Table 1). Histological results (Figure 3) were compared to facilitate the identification of the optimal perfusion condition.
Selection of perfu...

Watch this Scientific Journal Video about Normothermic Ex Vivo Liver Machine Perfusion in Mouse at JoVE.com

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

A normothermic ex vivo liver perfusion (NEVLP) system was created for mouse livers. This system requires experience in microsurgery but allows for reproducible perfusion results. The ability to utilize mouse livers facilitates the investigation of molecular pathways to identify novel perfusate additives and enables the execution of experiments focused on organ repair.

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

This protocol presents an optimized erythrocytes-free NEVLP system using mouse livers. Ex vivo preservation of mouse livers was achieved by employing ...

normothermic-ex-vivo-liver-machine-perfusion-in-mouse

Universidad de Cartagena UDEC

Research

JoVE Journal

Engineering

2.4K Views.  Jena University Hospital. A normothermic ex vivo liver perfusion (NEVLP) system was created for mouse livers. This system requires experience in microsurgery but allows for reproducible perfusion results. The ability to utilize mouse livers facilitates the investigation of molecular pathways to identify novel perfusate additives and enables the execution of experiments focused on organ repair.

Video: Normothermic Ex Vivo Liver Machine Perfusion in Mouse

High-resolution, High-speed, Three-dimensional Video Imaging with Digital Fringe Projection Techniques

Digital fringe projection (DFP) techniques provide dense 3D measurements of dynamically changing surfaces.&#160;Like the human eyes and brain, DFP uses triangulation between matching points in two views of the same scene at different angles to compute depth.&#160;However, unlike a stereo-based method, DFP uses a digital video projector to replace one of the cameras1. The projector rapidly projects a known sinusoidal pattern onto the subject, and the surface of the subject distorts these patterns in the camera&#8217;s field of view. Three distorted patterns (fringe images) from the camera can be used to compute the depth using triangulation.
Unlike other 3D measurement methods, DFP techniques lead to systems that tend to be faster, lower in equipment cost, more flexible, and easier to develop.&#160;DFP systems can also achieve the same measurement resolution as the camera.&#160;For this reason, DFP and other digital structured light techniques have recently been the focus of intense research (as summarized in1-5). Taking advantage of DFP, the graphics processing unit, and optimized algorithms, we have developed a system capable of 30&#160;Hz 3D video data acquisition, reconstruction, and display for over 300,000 measurement points per frame6,7.&#160;Binary defocusing DFP methods can achieve even greater speeds8.
Diverse applications can benefit from DFP techniques. Our collaborators have used our systems for facial function analysis9, facial animation10, cardiac mechanics studies11, and fluid surface measurements, but many other potential applications exist.&#160;This video will teach the fundamentals of DFP techniques and illustrate the design and operation of a binary defocusing DFP system.

This video describes the fundamentals of digital fringe projection techniques, which provide dense 3D measurements of dynamically changing surfaces.&#160;It also demonstrates the design and operation of a high-speed binary defocusing system based on these techniques.

Digital fringe projection (DFP) techniques provide dense 3D measurements of dynamically changing surfaces.&#160;Like the human eyes and brain, DFP uses ...

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.

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.

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 ...

Medicine

Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

Metabolic diseases such as diabetes, pre-diabetes, non-alcoholic fatty liver disease (NAFLD), and nonalcoholic steatohepatitis (NASH) are becoming increasingly common. Ex vivo liver perfusions allow for a comprehensive analysis of liver metabolism using nuclear magnetic resonance (NMR), in nutritional conditions that can be rigorously controlled. As in silico simulations remain a primarily theoretical means of assessing hormone actions and the effects of pharmaceutical intervention, the perfused liver remains one of the most valuable test beds for understanding hepatic metabolism. As these studies guide basic insights into hepatic physiology, results must be accurate and reproducible. The greatest factor in the reproducibility of ex vivo hepatic perfusion is the quality of surgery. Therefore, we have introduced an organized and streamlined method to perform ex vivo mouse liver perfusions in the context of in situ NMR experiments. We also describe a unique application and discuss common issues encountered in these studies. The overall purpose is to provide an uncomplicated guide to a technique we have refined over several years that we deem the golden standard for obtaining reproducible results in hepatic resections and perfusions in the context of in situ NMR experiments. The distance to the center of the field for the magnet as well as the inaccessibility of the tissue to intervention during the NMR experiment makes our methods novel.

Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

The protocol describes a straightforward method of resectioning an intact mouse liver for metabolic studies through portal vein perfusion.

Metabolic diseases such as diabetes, pre-diabetes, non-alcoholic fatty liver disease (NAFLD), and nonalcoholic steatohepatitis (NASH) are becoming ...

Biochemistry

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 ...

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, ...

Orthotopic Pig Liver Transplant Preservation Using Machine Perfusion

Preservation of Porcine Donation after Circulatory Death (DCD) Liver by Perfusion and Orthotopic Liver Transplantation

Conventional static cold storage (SCS) exacerbates ischemic injury in the DCD liver, leading to severe complications for transplant recipients. To address this issue, clinical application of MP technology for donor liver preservation is underway. Simultaneously, efforts are focused on the development of various MP instruments, validated through relevant animal model experiments. Effective large animal trials play a pivotal role in clinical applications. However, challenges persist in the ex vivo preservation of DCD livers and the transplantation procedure in pigs. These hurdles encompass addressing the prolonged preservation of donor livers, conducting viability tests, alleviating ischemic injuries, and shortening the anhepatic phase. The use of a variable temperature-controlled MP device facilitates the prolonged preservation of DCD livers through sequential Dual Hypothermic Oxygenated Machine Perfusion (DHOPE) and Normothermic Machine Perfusion (NMP) modes. This protocol enhances the porcine OLTx model by improving the quality of DCD livers, optimizing the anastomosis technique, and reducing the duration of the anhepatic phase.

Author Spotlight: Advancements and Challenges in Machine Perfusion for Liver Transplantation

This protocol outlines the methodology of establishing a porcine model utilizing variable temperature-controlled Machine Perfusion (MP) for the preservation of the donor liver, followed by orthotopic liver transplantation (OLTx). It aims to promote the success rate of OLTx using donor donation after circulatory death (DCD) liver and establish a stable model.

Conventional static cold storage (SCS) exacerbates ischemic injury in the DCD liver, leading to severe complications for transplant recipients. To address ...

Bioengineering

Avoiding Ischemia Reperfusion Injury in Liver Transplantation

Currently, ex situ machine perfusion is a burgeoning technique that provides a better preservation method for donor organs than conventional static cold preservation (0&#8211;4 &#176;C). A continuous blood supply to organs using machine perfusion from procurement and preservation to implantation facilitates complete prevention of ischemia reperfusion injury and permits ex situ functional assessment of donor livers before transplantation. In this manuscript, we provide a step-by-step ischemia-free liver transplantation protocol in which an ex situ normothermic machine perfusion apparatus is used for pulsatile perfusion through the hepatic artery and continuous perfusion of the portal vein from human donor livers to recipients. In the perfusion period, biochemical analysis of the perfusate is conducted to assess the metabolic activity of the liver, and a liver biopsy is also performed to evaluate the degree of injury. Ischemia-free liver transplantation is a promising method to avoid ischemia-reperfusion injury and may potentially increase the donor pool for transplantation.

Presented here is a protocol to provide a step-by-step ischemia-free liver transplantation protocol under ex situ normothermic machine perfusion (37 &#176;C) of human livers from donors to recipients.

Currently, ex situ machine perfusion is a burgeoning technique that provides a better preservation method for donor organs than conventional static cold ...

Development and Validation of a Porcine Ex Situ Isolated Liver Perfusion Model 

Porcine Normothermic Isolated Liver Perfusion

Porcine models of liver ex situ normothermic machine perfusion (NMP) are increasingly being used in transplant research. Contrary to rodents, porcine livers are anatomically and physiologically close to humans, with similar organ size and bile composition. NMP preserves the liver graft at near-to-physiological conditions by recirculating a warm, oxygenated, and nutrient-enriched red blood cell-based perfusate through the liver vasculature. NMP can be used to study ischemia-reperfusion injury, preserve a liver ex situ before transplantation, assess the liver's function prior to implantation, and provide a platform for organ repair and regeneration. Alternatively, NMP with a whole blood-based perfusate can be used to mimic transplantation. Nevertheless, this model is labor-intensive, technically challenging, and carries a high financial cost.
In this porcine NMP model, we use warm ischemic damaged livers (corresponding to donation after circulatory death). First, general anesthesia with mechanical ventilation is initiated, followed by the induction of warm ischemia by clamping the thoracic aorta for 60 min. Cannulas inserted in the abdominal aorta and portal vein allow flush-out of the liver with cold preservation solution. The flushed-out blood is washed with a cell saver to obtain concentrated red blood cells. Following hepatectomy, cannulas are inserted in the portal vein, hepatic artery, and infra-hepatic vena cava and connected to a closed perfusion circuit primed with a plasma expander and red blood cells. A hollow fiber oxygenator is included in the circuit and coupled to a heat exchanger to maintain a pO2 of 70-100 mmHg at 38 &#176;C. NMP is achieved by a continuous flow directly through the artery and via a venous reservoir through the portal vein. Flows, pressures, and blood gas values are continuously monitored. To evaluate the liver injury, perfusate and tissue are sampled at predefined time points; bile is collected via a cannula in the common bile duct.

The porcine model of liver normothermic machine perfusion (NMP), described here, can be successfully used to study NMP as a preservation strategy, a tool for viability assessment, and a platform for organ repair. It holds a high translational value, however it is technically challenging and labor-intensive.

Porcine models of liver ex situ normothermic machine perfusion (NMP) are increasingly being used in transplant research. Contrary to rodents, porcine ...

In Situ Mouse Liver Perfusion Model to Study Metabolic Responses to Substrates

Hepatic Glucose Production, Ureagenesis, and Lipolysis Quantified using the Perfused Mouse Liver Model

The liver has numerous functions, including nutrient metabolism. In contrast to other in vitro and in vivo models of liver research, the isolated perfused liver allows the study of liver biology and metabolism in the whole liver with an intact hepatic architecture, separated from the influence of extra-hepatic factors. Liver perfusions were originally developed for rats, but the method has been adapted to mice as well. Here we describe a protocol for in situ perfusion of the mouse liver. The liver is perfused antegradely through the portal vein with oxygenated Krebs-Henseleit bicarbonate buffer, and the output is collected from the suprahepatic inferior vena cava with clamping of the infrahepatic inferior vena cava to close the circuit. Using this method, the direct hepatic effects of a test compound can be evaluated with a detailed time resolution. Liver function and viability are stable for at least 3 h, allowing the inclusion of internal controls in the same experiment. The experimental possibilities using this model are numerous and may infer insight into liver physiology and liver diseases.

Author Spotlight: Investigating Hepatic Adaptations and Prediabetic Progression in Liver Diseases 

Here, we present a robust method for in situ perfusion of the mouse liver to study the acute and direct regulation of liver metabolism without disturbing the hepatic architecture but in the absence of extra-hepatic factors.

The liver has numerous functions, including nutrient metabolism. In contrast to other in vitro and in vivo models of liver research, the isolated perfused ...

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

Summary

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Chapters in this video

High-resolution, High-speed, Three-dimensional Video Imaging with Digital Fringe Projection Techniques

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

Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

Ex Situ Normothermic Machine Perfusion of Donor Livers

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

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

Preservation of Porcine Donation after Circulatory Death (DCD) Liver by Perfusion and Orthotopic Liver Transplantation

Avoiding Ischemia Reperfusion Injury in Liver Transplantation

Porcine Normothermic Isolated Liver Perfusion

Hepatic Glucose Production, Ureagenesis, and Lipolysis Quantified using the Perfused Mouse Liver Model