Liver Cold Storage and Transplantation in the Cold-Adaptive Daurian Ground Squirrels

Summary

Here, we present a protocol to establish a long-term liver cold storage and transplantation model in Daurian ground squirrel (DGS), which has been technically challenging in standard rodent models.

Abstract

In liver transplantation, the functional recovery of donor grafts following extended cold storage is a major challenge. The same obstacle applies to standard rodent models (e.g., mice and rats) for liver transplantation despite their well-established surgical protocols, thus restricting research on key topics such as post-transplantation functional recovery and host-graft immunocompatibility. Here, we successfully established a novel liver transplantation model using an obligate hibernator, the Daurian ground squirrel (DGS; Spermophilus dauricus), which demonstrates intrinsic adaptation to ultra profound hypothermia-rewarming and ischemia-reperfusion transitions. In our hands, transplantation of liver grafts following 24-h cold storage resulted in 100% survival of DGS recipients within the 7-day observation time frame, whilst the same procedure in Sprague-Dawley (SD) rats had 0% survival 24 h following the surgery. Thus, the DGS and other cold-adaptive mammalian models provide a unique and robust platform to investigate mechanisms underlying organ cold adaptation and transplantation tolerance, which can support the development of novel treatment or surgery strategies in the field of transplantation medicine.

Introduction

Cold-rewarming and ischemia-reperfusion are significant contributors to liver graft injury, increasing the risk of early allograft dysfunction. Although liver transplantation in standard rodent organisms such as mice and rats has been successfully established^1,2, because studies on these cold-intolerant species provide limited insights into the mechanisms of cold adaptation, advancements in organ preservation and transplantation medicine have been hindered.

Developing transplantation research models in suitable cold-adaptive species may fill such a gap. For example, mammalian hibernators have evolved to survive lengthy periods of low ambient temperature and/or food scarcity^3,4. During hibernation, their physiology transitions in cycles of torpid and interbout arousal states so that they are well adapted to maintaining a low core body temperature and enduring rapid rewarming and blood reperfusion. Remarkably, ground squirrels that can hibernate demonstrate more resilience to ischemia-reperfusion injuries even in their active, euthermic state than rats^5,6. In recent years, hibernator-inspired studies have unveiled some cold-adapting mechanisms and druggable targets that can be modulated to significantly alleviate cold storage-induced injuries and improve functional recovery of isolated tissues and organs from non-hibernating species^7,8,9,10,11.

Adapted from protocols developed for liver transplantation in rats^12,13,14, here we introduce a protocol for liver procurement, static cold storage, and transplantation in Daurian ground squirrel (DGS; Spermophilus dauricus), a hibernating rodent species that resides in Northern China. In our hands, DGS graft recipients had 100% post-transplantation survival with grafts from either euthermic (active) or hypothermic (torpid) donors stored at 4 ˚C in University of Wisconsin (UW) solution for 24 h. In stark contrast, the survival rate in Sprague-Dawley (SD) rats subjected to the same workflow was 0%, clearly showing the advantage of the DGS liver transplantation model. We expect long-term cold storage and transplantation of other organs in DGS or other lab-bred cold-adaptive species, such as the 13-lined ground squirrel (Ictidomys tridecemlineatus) and the Syrian hamster (Mesocricetus auratus), can also be achieved, thus providing unique opportunities for researchers to study long-term graft functional recovery and host-graft compatibility, and test new medicine.

Protocol

All animal experiments were approved by the Peking University Institutional Committee for Animal Care and Use (LSC-WangSQ-2). The appropriate instruments used for the study are listed in the Table of Materials and depicted in Figure 1.

NOTE: Handling and breeding of wild-caught hibernating rodent species, such as the 13-lined ground squirrels, in a laboratory environment has been described^15,16. The DGSs used in this study were produced in the laboratory by female pregnant DGSs captured in the grassland of Zhangjiakou city. They were about 1-2 years old at the time of the experiment. As for the original pregnant female DGSs, parasites had been expelled before the animals arrived at the laboratory. Quarantine and inspection for common viruses were conducted upon the arrival of the animals, and those with negative test results were bred in the facility.

1. Preoperative preparation

1. Experimental animals
   1. Use adult DGS, 1-2 years old, both male and female, weighing between 180 g and 400 g as donors and recipients. The recipient's weight was slightly greater than the donor's, with a difference of less than 50 g.
   2. Use adult SD rats, 8-10 weeks, both male and female, weighing 200-300 g, as donors and recipients in the control group. During the whole experimental procedure, wear surgical masks, lab coats, and nitrile gloves, as well as extra heavy leather gloves during the handling and transfer of DGS animals.
      NOTE: The surgical techniques for rats have been well documented elsewhere. The protocol described here focuses on the liver transplantation techniques in DGS.
2. Make cuffs for the portal vein (PV) and infrahepatic inferior vena cava (IHIVC) using 6 F and 8 F introducer sheaths, respectively. Each cuff comprised a main body and extensions measuring 2 mm in length. Create multiple grooves around the circumference of the cuff by gently clamping with mosquito forceps (Figure 2).
   NOTE: The diameters of the PV and IHIVC vary significantly depending on the size and sex of the DGS. Therefore, it is advisable to prepare 5-F, 6-F, 7-F, and 8-F cuffs preoperatively for appropriate selection. Typically, a 5-F or 6-F cuff is suitable for the PV, while a 7-F or 8-F cuff is appropriate for the IHIVC.
3. Make stents for the bile duct (3 mm) and hepatic artery (2 mm) by cutting 22 G and 24 G intravenous catheters into double-sided slanted tubes using a scalpel.
   NOTE: Avoid applying excessive pressure to the stents, as this may lead to narrowing or occlusion of the lumen, compromising recipient survival.

2. Donor operation

1. Initial steps
   1. Use 5 % isoflurane concentration at an airflow rate of 1 L/min to induce anesthesia and 2 % at 1 L/min for maintenance.
   2. Position the DGS in a supine position on a board with a heating pad to maintain body temperature intraoperatively. Shave the fur from the abdominal area and disinfect the skin with 3 alternating applications of povidone-iodine and 70% ethanol.
2. Liver isolation prior to perfusion
   1. Confirm the depth of anesthesia via a toe-pinch. Make a cruciate incision extending from the xiphoid process to the pubic symphysis and laterally to the mid-axillary lines. Retract the xiphoid process using mosquito forceps and place a 5 mL syringe beneath its back as support.
   2. Transect the falciform ligament. Isolate the left diaphragmatic vein from the suprahepatic inferior vena cava (SHIVC) and ligate it with a 7-0 suture. Cauterize the hepato-splenic ligament.
   3. Carefully free the IHIVC above the right renal vein from surrounding tissues using cotton swabs. Ligate the right adrenal and lumbar veins draining into the IHIVC near the liver with 7-0 sutures.
   4. Make a small incision on the anterior wall of the common bile duct (CBD). Insert the bile duct stent and secure it with a 7-0 silk suture. Leave one end of the suture approximately 5 mm in length to facilitate later anastomosis with the recipient's CBD.
   5. Isolate the proper hepatic artery (PHA), gastroduodenal artery (GDA), and common hepatic artery (CHA) from surrounding tissues.
   6. Administer 1.5 mL of heparinized saline (300 IU/mL) into the penile vein (male donors) or the IHIVC (female donors).
3. Liver perfusion
   1. Isolate the abdominal aorta below the left renal vein. Insert a 7 G intravenous needle into the aorta and secure it with a clamp.
   2. Incise the diaphragm and place a vascular clamp on the thoracic aorta.
   3. Perfuse the liver via the aortic cannula with 20-30 mL of cold UW solution at a rate of 180-210 mL/h using an infusion pump. Incise the intrathoracic vena cava and the IHIVC above the right renal vein to allow efflux of the perfusate.
      NOTE: Adjust the position of liver lobes using cotton swabs to ensure uniform perfusion, paying particular attention to the middle and right lobes. Perfusion is complete when the entire liver exhibits a uniform pallor.
   4. During perfusion, excise the gallbladder.
      NOTE: Unlike rats, DGS possess a gallbladder. Removing the gallbladder and flushing the bile ducts with UW solution are essential to prevent postoperative biliary obstruction due to bile coagulation.
4. Liver graft harvest post-perfusion
   1. Using micro-forceps, isolate the PV from the surrounding connective tissue. Ligate and transect the pyloric vein with a 7-0 suture. Transect the PV trunk at the level of the splenic vein.
   2. Ligate and transect the GDA distally. Create a small incision on the anterior wall of the CHA. Insert the hepatic artery stent into the CHA and secure it with a 7-0 silk suture, leaving one end approximately 5 mm long to facilitate later anastomosis with the recipient's CHA.
   3. Dissect the SHIVC as close to the diaphragm as feasible.
   4. Dissect the attachments between the caudate lobe and the stomach. After severing the dorsal ligaments, fully mobilize the liver.
   5. Place the liver into a 7-cm steel bowl containing cold UW solution. Transfer the bowl to a 4 °C refrigerator for preservation.
      NOTE: For liver cold preservation exceeding 24 h, add antibiotics to the UW solution to reduce the risk of infection; consider replacing the UW solution every 24 h.

3. Ex vivo graft preparation

NOTE: Prepare an open ice container. The whole back table preparation was done on ice.

1. Secure the handle of the PV cuff with a vascular clamp and stabilize it using mosquito forceps. Gently thread the PV through the cuff, ensuring no twisting occurs. Fold the PV over the cuff and secure it with a 7-0 suture (Figure 3).
2. Repeat the above procedure (step 3.1) for the IHIVC cuff.
3. Perfuse the bile duct via the cystic duct with 2 mL of UW solution. Subsequently, ligate the cystic duct.
4. Flush the liver graft with 8-10 mL of cold saline through the PV cuff to remove residual UW solution after cold storage.
5. Place a single 7-0 silk suture around the IHIVC between the attached cuff and the right inferior lobe of the liver.
6. Rotate the liver to expose its superior aspect. Place 8-0 stay sutures at both lateral corners of the SHIVC, inserting from the exterior to the interior.

4. Recipient operation

1. For initial steps, refer to the donor operation protocol and repeat step 2.1.
2. Recipient liver isolation
   1. Confirm the depth of anesthesia via a toe-pinch. Perform a midline incision from the xiphoid process to the pubic symphysis. Retract the xiphoid process cranially using mosquito forceps and position a 5 mL syringe beneath to elevate the SHIVC. Laterally retract the bilateral costal arches.
   2. Perform ligament and vessel management as described in step 2.2.2.
   3. Isolate IHIVC as described in step 2.2.3.
   4. Sever the ligamentous attachments between the liver and retroperitoneum. Carefully pass a blunt forceps behind the SHIVC and place a 7-0 silk suture around it.
   5. Ligate the CBD proximally with a 7-0 silk suture and transect it above the ligature. Use a cotton tip to gently separate the bile duct from the PV.
   6. Transect the GDA and PHA between ligations. Create a Y-shaped bifurcation at the end of the CHA. Leave a 2 cm length of suture on the PHA ligature for later use.
   7. Free the right and left branches of the PV. Place 5 cm, 7-0 stay sutures on each branch for subsequent manipulation.
3. Recipient liver removal
   1. Reduce the isoflurane concentration to 0.5% with a 0.5 L/min airflow.
      NOTE: Maintain a respiratory rate of approximately one breath per second by adjusting the isoflurane vaporizer. Continuously assess anesthesia depth throughout the anhepatic phase to prevent intraoperative awareness or excessive anesthesia leading to respiratory depression.
   2. Clamp the IHIVC above the right renal vein, followed by clamping the PV above the pyloric vein. Inject 1.5 mL of saline into the right PV in 10 s using a 2 mL syringe to flush blood from the liver.
   3. Gently retract the diaphragm by pulling the 7-0 silk suture placed around the SHIVC, then apply a bulldog clamp to the SHIVC, including a portion of the diaphragm.
   4. Transect the SHIVC just above the liver. Secure the 7-0 stay sutures on both PV branches and transect the PV above these ligatures. Transect the IHIVC close to the liver parenchyma. Promptly remove the liver by severing any remaining ligamentous attachments.
4. SHIVC reconstruction
   1. Position the liver graft orthotopically within the abdominal cavity. Utilize the attached 8-0 sutures to place stay sutures at both corners of the recipient's SHIVC, inserting from the interior to the exterior, and secure with knots.
   2. Gently traction the loose end of the right suture laterally using a vascular clip to align the SHIVC.
   3. Suture the posterior wall with 8-10 continuous stitches, beginning at the left corner. Proceed to close the anterior wall with 10-12 stitches, moving from right to left using the same suture. Prior to completing the anterior wall closure, flush the SHIVC cavity to expel any air and prevent air embolism (Figure 4).
5. PV reconstruction
   1. Elevate the PV using the 5 cm stay sutures placed on both bifurcations, applying gentle upward traction with vessel clamps. Make an incision at the midpoint of the PV branches.
   2. Flush the PV lumen with heparinized saline (300 IU/mL), and then insert the cuff into the PV and secure it with a 7-0 silk suture.
   3. Sequentially release the clamps on the PV and SHIVC to reestablish graft perfusion and end the anhepatic phase (Figure 5).
6. Perform the IHIVC anastomosis similarly to the PV procedure. Release the 7-0 suture around the donor side first, then remove the clamp from the recipient side.
7. Hepatic artery reconstruction
   1. Apply a microvascular clip to occlude blood flow in the recipient's CHA.
   2. Create a small incision at the bifurcation of the Y-configuration at the terminus of the recipient's CHA. Flush the CHA lumen with heparinized saline (300 IU/mL), and then insert the stent into the CHA and secure it with a 7-0 silk suture.
   3. Tie the stay sutures on both donor and recipient sides to prevent stent displacement. Release the microvascular clip to restore arterial flow (Figure 6).
8. Bile duct reconstruction:
   1. Make a small incision on the anterior wall of the recipient's CBD. Insert the bile duct stent into the lumen of the recipient's CBD and secure it with a 7-0 silk suture.
   2. Tie the stay sutures on both donor and recipient sides to prevent stent displacement (Figure 7).
9. Close the abdominal incision in two layers using continuous running 3-0 sutures after the completion of the reconstruction procedures (Figure 8).
10. Place the recipient in a clean, heated cage, and monitor recovery every 1-2 h before returning the animal to the animal housing facility. Subcutaneously inject buprenorphine (0.3 mg/kg) once a day for 3 days for post-operative analgesia.

Results

The mean durations for the surgical procedures were as follows: donor surgery averaged 24.8 ± 2.3 min, back table operation 15.4 ± 1.3 min, recipient surgery 52.7 ± 5.1 min, and the anhepatic phase 14.2 ± 1.5 min.

Recipients of liver grafts preserved in cold UW solution for 24 h exhibited a 100% survival rate at 7 days post-transplantation, regardless of whether they were in a hibernation state prior to transplantation (n = 8) o...

Discussion

Building upon Kamada's double-cuff technique², we have successfully established the first orthotopic liver transplantation model in DGS. Due to anatomical differences between the DGS and traditional rat models, specific considerations in material preparation, surgical procedures, and postoperative management are essential.

Factors such as hibernation and fat reserves contribute to significant individual weight variations in adult DGS, ranging from 150 g to 400 g, with n...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
3-0 sutures with needles	Jinhuan Medical Products Co., Ltd	R315	For skin closure
7-0 silk suture	Braintree, sci, inc	SUT-S103
7F Introducer Sheath	Beijing Demax Medical Technology Co., Ltd.	DQ07083818S	Cuff for IHIVC
8-0 sutures with needles	Jinhuan Medical Products Co., Ltd.	H821	For SHIVC anastomsis
8F Introducer Sheath	Beijing Demax Medical Technology Co., Ltd.	DQ08083818S	Cuff for IHIVC
Anesthesia Machine	RWD life Science Co.	R500
Blood Stop Cautery	Ningbo Shunye Medical Co., Ltd.	BDD-YE-DT-1
Bulldog clamp (slightly curved)	Jinzhong Medical Products Co., Ltd.	XEC120	SHIVC
Buprenorphine	Tianjin Institute of Pharmaceutical Research Pharmaceutical Co., Ltd	H12020275	Painkiller
Cefuroxim sodium	Esseti FarmaceuticiS.r.l	H20160013	Antibiotic
Cotton swabs	Fuqing Health & Integral Medical	20230R
Curved micro forceps	RWD life Science Co.	F11031-11	Surgical tool
Curved micro scissors	66 Vision Tech Co., Ltd.	54108B	Surgical tool
Gauze swabs	Yubei Medical Materials Co., LTD	21080274
Heparin sodium injection	North China Pharmaceutical Co., LTD	2101131-2
Infusion Pump	Howkmed Co., Ltd.	HK-400
Isoflurane	RWD life Science Co.	21070201
Micro needle holder	World Precision Instruments Co., Ltd.	555408NT	Surgical tool
Micro vessel clip	RWD life Science Co.	R31005-06	PV and IHIVC
Micro Vessel clip applicator	RWD life Science Co.	R34001-14	Surgical tool
Needle holder	RWD life Science Co.	F31025-13	Abdominal closure
Saline	Zhejiang Tianrui Pharmaceutical Co.,Ltd	716092103
Straight micro forceps	RWD life Science Co.	F11029-11	Surgical tool
Surgical microscope	Leica	M650
Surgical platform	N/A	N/A	Custom, magnetic
Tissue forceps	RWD life Science Co.	F12002-12	Surgical tool
Tissue scissor	RWD life Science Co.	S12003-09	Surgical tool
UW solution	Organ Recovery Systems	SPS-1	Organ perfustion and storage