Normothermic Ex Vivo Liver Machine Perfusion in Mouse

Summary

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.

Abstract

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 °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'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.

Introduction

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 underway^3,4.

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 recognized^6,7. 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'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 "organ repair."

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 "normothermic liver perfusion," "ex vivo or in vitro," and "rats". 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 °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.

Protocol

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 Thüringer Landesamt für Verbraucherschutz, Thuringia, Germany (Approval-Number: UKJ - 17 - 106).

NOTE: Male C57BL/6J mice weighing 34 ± 4 g (mean ± standard error of the mean [SEM]) were used as liver donors. They were maintained under controlled environmental conditions (50% humidity and 18 - 23 °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 °C.

1. Preparation

1. Setting up the operation table
   1. Autoclave all surgical instruments and consumables for sterilization purposes.
   2. Turn on all equipment, including the warming board and electrocoagulation.
   3. Place one 50 mL syringe with 25 mL heparinized (2,500 U/L) saline in a warm incubator (37 °C).
   4. 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.
   5. 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.
   6. 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.
2. Self-made portal vein cannula
   1. 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.
   2. Trim the tip of the cannula resulting in a sharp triangle.
3. Preparation of heparinized saline
   1. Prepare 25 mL of heparinized saline with a final concentration of 2,500 IU/mL.
   2. Remove all air bubbles and place the syringe in the 40 °C incubator.
4. Demonstration of the perfusion system
   1. See Figure 1 for the main components of the machine perfusion system.
5. Set up of the organ chamber
   1. See Figure 2 for the layout of the organ chamber.
6. Set up of the perfusion system
   1. Turn on the lab chart program for pressure monitoring.
   2. Connect the pressure calibrator and pressure sensor at the organ chamber level.
   3. Adjust the pressure calibrator to read 0 mmHg and check the corresponding value on the pressure control software.
   4. Adjust the pressure calibrator to read 20 mmHg and again check the corresponding value on the pressure control software.
   5. Turn on the water bath, and prewarm the organ chamber to 40 °C.
   6. Flush the entire plumbing system twice with distilled deionized water for 30 min each, ensuring complete removal of the sanitizing solution.
   7. Initiate the circulation of the disinfection solution throughout the entire system for a duration of 20 min to ensure thorough disinfection.
   8. Turn on the gas mixture (95% oxygen (O₂) and 5% carbon dioxide (CO₂).
7. Perfusate filling
   1. Supplement 250 mL of Williams' 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' E medium.
   2. 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.
   3. Turn on the peristaltic pump at medium speed (15 mL/min) to prime the perfusion system with the oxygenated medium.

2. Liver explantation

1. Pre-surgery preparation
   1. Weigh the animal. Prepare the analgesic buprenorphine (0.3 mg/mL) (0.05 mg/kg body weight).
   2. Connect the induction chamber with the wall socket. Turn oxygen to 0.5 L/min. Turn isoflurane to 3%.
   3. Place the animal in the chamber until deep anesthesia (righting reflex positive) is reached.
   4. Use a micro syringe to apply body weight-adapted dose of analgesia subcutaneously.
   5. Use an electric shaver to trim the fur on the abdominal skin.
   6. 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.
2. Preparation of the mouse abdomen
   1. Place the mouse in a supine position.
   2. Test interdigital reflex to double confirm the appropriate depth of anesthesia. Fix all four limbs with tape.
   3. 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.
   4. 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.
   5. Extend the skin incision bilaterally to the midaxillary line on both sides.
   6. Carefully make a 2 cm longitudinal incision along the linea alba using spring scissors.
   7. Cut through the abdominal muscle layer with electrocoagulation and Vannas spring scissors.
   8. Carefully place a piece of wet gauze to protect the liver from electrocoagulation.
   9. Use a 6 - 0 silk suture with the round needle to retract the xiphoid process for better exposure of the coronary ligament.
   10. Use two rib retractors to fully expose the abdominal cavity of the mouse.
   11. Carefully move the small intestine out of the abdominal cavity with a wet cotton swab to fully expose the hilum.
3. Common bile duct preparation
   1. Transect the falciform, phrenic, and gastrohepatic ligaments with sharp scissors.
   2. Carefully free the common bile duct using fine curved forceps without teeth.
      ​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.
   3. Place two 6 - 0 silk suture loops over the common bile duct in preparation for the next step.
4. Common bile duct cannulation
   1. 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.
   2. Use vessel cannulation forceps to grasp the bile duct cannula and push it into the bile duct.
   3. 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.
   4. Observe the bile flow in the cannula after successful cannulation.
5. Portal vein preparation
   1. 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.
   2. 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.
   3. Place the second suture loop for later fixation of the PV as close to the hepatic hilum as possible.
6. Portal vein cannulation
   1. Use an arterial clip to close the distal portal vein.
   2. 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.
   3. Secure the PV cannula with the pre-placed 6 - 0 suture loop.
7. Liver flushing
   1. Increase isoflurane to 5% and euthanize the mouse with an overdose of isoflurane inhalation.
   2. Take prewarmed heparin saline solution from the incubator. Remove all air bubbles formed inside the heparinized saline.
   3. Fix the syringe with prewarmed heparinized saline into the syringe pump.
   4. 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.
   5. Observe the color of the liver at the end of the flushing procedure. Excise the liver once the color turns to a homogenous yellow.
   6. Transect the diaphragm, suprahepatic inferior vena cava, infra hepatic vena cava, hepatic artery, distal portal vein, and any remaining connective tissue.
   7. Place the liver into the Petri dish.

3. Liver and chamber connection

1. Liver transfer
   1. Carefully transfer the liver into the organ chamber using a Petri dish.
   2. 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.
2. Portal vein cannula connection
   1. Slowly infuse normal saline into the portal vein cannula with a syringe to evacuate the air bubbles in the cannula.
   2. Connect the portal vein cannula into the perfusate outflow tube in the organ chamber.
3. Bile duct cannula connection
   1. Guide the mouse bile duct cannula through the valve of a rubber cap which is connected to the organ chamber.
   2. Insert the bile duct cannula into a pre-prepared 0.5 mL microtube with a small hole in the lid.
   3. Place the microtube on clay outside of the organ chamber.

4. Adjust the flow rate according to PV pressure

1. Turn on the peristaltic pump from 1 mL/min.
2. Check the portal vein pressure reading to adjust the flow rate.
3. 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.

5. Sample collection

1. Obtain inlet perfusate samples from the portal vein inflow tube and outlet perfusate samples from the organ chamber in 3h intervals.
2. Collect samples from all liver lobes for histological analysis at the end of the perfusion period of 12 h.

Results

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...

Discussion

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...

Materials

Name	Company	Catalog Number	Comments
0.5 ml Micro Tube PP	Sarstedt	72699
1 Fr Rubber Cannula	Vygon	Sample Cannula
10 µL Micro Syringe	Hamilton	701N
2 Fr Rubber Cannula	Vygon	Sample Cannula
24 G Butterfly Cannula	Terumo	SR+OF2419
26 G Butterfly Cannula	Terumo	SR+DU2619WX
30 G Hypodermic Needle	Sterican	100246
50 ml Syringe Pump	Braun	110356
6-0 Perma-Hand Seide	Ethicon	639H
Arterial Clip	Braun	BH014R
Autoclavable Moist Chamber	Hugo Sachs Elektronik	73-4733
Big Cotton Applicator	NOBA Verbandmittel Danz GmbH	974018
Bubble Trap	Hugo-Sachs-Elektronik	V83163
Buprenovet (0.3 mg / ml)	Elanco	/
CIDEX OPA solution (2 L)	Cilag GmbH	20391
Electrosurgical Unit for Monopolar Cutting VIO® 50 C	ERBE	/
Fetal Bovine Serum(500 ml)	Sigma-Aldrich	F7524-500ML
Gas Mixture (95 % oxygen & 5 % carbon dioxide)	House Supply	/
Heating Circulating Baths	Harvard-Apparatus	75-0310
Heparin 5000 (I.E. /5 ml)	Braun	1708.00.00
Hydrocortisone (100 mg / 2 ml)	Pfizer	15427276
Insulin(100 IE / ml)	Sigma	I0516-5ML
Iris Scissors	Fine Science Instruments	15000-03
Isofluran (250 ml)	Cp-Pharma	1214
Membrane Oxygenator	Hugo Sachs Elektronik	T18728
Microsurgery Microscope	Leica	M60
Mouse Retractor Set	Carfil Quality	180000056
NanoZoomer 2.0 HT	Hamamatsu	/
Non-Woven Sponges	Kompressen	866110
Penicillin Streptomycin (1 mg / ml)	C.C.Pro	Z-13-M
Perfusion Extension Tube (30 cm)	Braun	4256000
Peristaltic Pump	Harvard-Apparatus	P-70
Petri Dishc 100x15 mm	VWR®	391-0578
Povidon-Jod (Vet-Sep Spray)	Livisto	799-416
Pressure Transducer Simulator	UTAH Medical Products	650-950
Reusable Blood Pressure Transducers	AD Instruments	MLT-0380/D
S & T Vessel Cannulation Forceps	Fine Science Instruments	00608-11
Small Cotton Applicator	NOBA Verbandmittel Danz GmbH	974116
Straight Forceps 10 cm	Fine Science Instruments	00632-11
Suture Tying Forceps	Fine Science Instruments	11063-07
Syringe 50ml Original Perfusor	Braun	8728810F-06
UT - 03 Cannula	Unique Medical, Japan	/
Vannas Spring Scissors	Fine Science Instruments	15018-10
Veterinary Saline (500 ml)	WDT	18X1807
Water Jacketed Reservoir  2 L	Harvard-Apparatus	73-3441
William's E Medium (500 ML)	Thermofischer Scientific	A1217601