Heterotopic Auxiliary Whole Liver Rat Transplant Model Utilizing a Hepaticoureterostomy for Allograft Rejection Studies

Summary

The rat heterotopic auxiliary liver transplant protocol described here offers a practical investigational tool for exploring mechanisms of hepatic allograft rejection. This model helps to alleviate the surgical hurdles and animal stress of orthotopic liver transplantation in rats.

Abstract

Small animal transplant models are indispensable for organ tolerance studies investigating feasible therapeutic interventions in preclinical studies. Rat liver transplantation (LTx) protocols typically use an orthotopic model where the recipients' native liver is removed and replaced with a donor liver. This technically demanding surgical procedure requires advanced micro-surgical skills and is further complicated by lengthy anhepatic and lower body ischemia times. This prompted the development of a less complicated heterotopic method that can be performed faster with no anhepatic or lower body ischemia time, reducing post-surgery stress for the recipient animal.

This heterotopic LTx protocol includes two main steps: excising the liver from the donor rat and transplanting the whole liver into the recipient rat. During the excision of the donor liver, the surgeon ligates the supra-hepatic vena cava (SHVC) and hepatic artery (HA). On the recipient side, the surgeon removes the left kidney and positions the donor liver with the portal vein (PV), infra-hepatic vena cava (IHVC), and bile duct facing the renal vessels. Further, the surgeon anastomoses the recipient's renal vein end to end with the IHVC of the liver and arterializes the PV with the renal artery using a stent. A hepaticoureterostomy is utilized for biliary drainage by anastomosing the bile duct to the recipient's ureter, permitting the discharge of bile via the bladder.

The average duration of the transplantation was 130 min, cold ischemia duration was around 35 min, and warm ischemia duration was less than 25 min. Hematoxylin and eosin histology of the auxiliary liver from syngeneic transplants showed normal hepatocyte structure with no significant parenchymal alterations 30 days post-transplant. In contrast, 8-day post-transplant allogeneic graft specimens demonstrated extensive lymphocytic infiltration with a Banff Schema rejection activity index score of 9. Therefore, this LTx method facilitates a low morbidity rejection model alternative to orthotopic LTx.

Introduction

Small animal LTx is an invaluable model for investigating mechanisms of liver rejection. Heterotopic auxiliary liver transplantation with portal vein arterialization (HALT-PVA) in rats was introduced in 1968 by Lee and Edgington¹when they reported using a recipient's renal vein and artery to re-vascularize a grafted auxiliary liver. Subsequently, Hess et al.² enhanced the protocol with the mitigation of functional competition between the native and auxiliary livers by reducing the native and donor liver size along with reconstructing the donor bile duct connection, resulting in long-term graft survival. Further refinements were made with the introduction of cuff anastomosis^3,4, and Schleimer et al.⁵ determined the optimal stent diameter for regulating blood flow to obtain physiological portal flow and avoid hyper- or hypo-perfusion of the graft. Other investigators developed significant alterations to the method by using the splenic⁶ or common iliac⁷ artery for graft blood supply, while some developed models that used only venous blood⁸ or only arterial blood via the hepatic artery⁹ to supply the auxiliary liver graft.

The present study hypothesized that functional competition from the native liver would not interfere with allograft rejection, so we developed a protocol based on the flow-regulated Schleimer model¹⁰ that did not include any size reduction of the native or auxiliary liver. The left side of the recipient was selected to locate the graft because it provided optimal orientation between the recipient's renal and donor liver vessels. Initially, we attempted biliary reconstruction via hepaticoduodenostomy but these trials simply confirmed Schleimer's assertion that "biliary drainage is the Achilles heel of liver transplantation"¹⁰. This prompted the development of a new technique where the bile duct is anastomosed end-to-end using a stent with the recipients' ureter, permitting the discharge of bile via the bladder. A noteworthy benefit of using a hepaticoureterostomy is that graft liver functionality can be monitored daily by observing the urine; a bile-producing liver graft colors the urine a bright yellow. Figure 1 represents the schematic overview of the HALT-PVA method.

An important advantage of heterotopic over orthotopic rat LTx relates to the absence of any anhepatic or total lower body ischemia time, which permits quicker and easier recoveries for heterotopic recipients. Additionally, LTx immunological studies utilizing orthotopic methods often rely on severe rejection or death of the recipient as an experimental endpoint, which is not the case with heterotopic transplants, where the animal remains healthy even if the allograft stops functioning due to rejection. Both of these features of the heterotopic method support principles of the international 3R's initiative (Replacement, Reduction, and Refinement)¹¹, which promotes a framework for minimizing the pain, suffering, and distress experienced by research animals and improving their welfare.

The HALT-PVA model reported here is a practical and reliable method for investigating the mechanisms of hepatic allograft rejection in preclinical studies. This useful experimental technique helps overcome the considerable surgical demands and animal stress of orthotopic LTx in rats. In the future, we intend to use this method to investigate the mechanisms of acute immune rejection while exploring novel targets and therapeutic strategies to suppress hepatic allograft rejection.

Protocol

Animals were bred and housed in specific pathogen-free conditions in the animal care facilities at the University of Wisconsin (UW)-Madison Institute for Medical Research in accordance with institutional guidelines. The study protocol (No. M006022) was approved by the Institutional Animal Care and Use Committee at the UW School of Medicine and Public Health, and all animals were treated ethically.

1. Animals

1. Use adult Lewis female rats weighing 205-235 g and Lewis males weighing 250-280 g as donor rats. Use adult male Lewis and Brown Norway rats weighing 365-420 g as recipients.
2. Perform syngeneic transplants by transplanting Lewis donors into Lewis recipients, while the allogeneic transplants utilized Lewis donors transplanted into Brown Norway recipients.
3. Perform all surgeries by two persons using a dual-head microscope.

2. Auxiliary liver donor procurement procedure

1. Anesthetize the donor rat with 5% isoflurane inhalation in an induction chamber. Record the weight of the rat and shave the abdomen with an electric clipper.
2. Position the rat in a supine position on a heated surgery pad with its nose in an anesthesia cone and immobilize the limbs with tape. Disinfect the abdomen with 75% alcohol and lower the isoflurane to 2%.
3. Make a longitudinal midline skin and muscle incision from the pubis to the xiphoid using scissors. Near the mid-point of the longitudinal incision, extend it laterally to the left and right, then install retractors on both sides of the abdominal wall and the xiphoid process.
4. Using moist cotton swabs, retract the intestines to the left side of the abdomen while using spring scissors to cut the gastro ligaments attached to the liver, then immobilize the intestines under moistened gauze. Cover the liver with a small piece of sterile gauze moistened with warm saline.
5. Use moist cotton swabs to retract the liver and dissect the falciform, triangular, hepatogastric, and hepatoduodenal ligaments. Next, cauterize with bipolar forceps and divide the para-esophageal vessels between the left lateral and anterior caudate lobe.
6. Using angle forceps or needle holders, dissect behind the SHVC inferior to the diaphragm, then pass a 5-0 silk suture under the SHVC and loosely tie a double knot for later use.
7. Retract the inferior right lateral lobe upward, cut the ligament, and immobilize under moistened gauze. Isolate the IHVC from the retroperitoneal tissue down to the right renal vein and ligate the right adrenal vein with a 6-0 silk suture as close to the IHVC as possible. Divide this vein later when the graft is removed.
8. Using a 27 G hydro-dissection needle (Figure 2A), dissociate the PV from surrounding connective tissue and separate it from the pyloric and splenic veins by ligating and dividing them using a 7-0 silk suture.
9. Isolate, ligate with 6-0 silk suture, and divide the common hepatic artery close to where it passes beneath the PV.
10. Ligate the bile duct with a 5-0 silk suture at the branching of the gastroduodenal artery while preserving the fat tissue around the bile duct as much as possible; in particular, avoid separating the bile duct from the proper hepatic artery while keeping the overall length as short as is practical.
11. Using spring scissors, make a small incision in the wall of the bile duct proximal to the ligation. Insert a 0.0215" diameter by 5 mm long polyimide tubing stent into the bile duct lumen and secure it with a 6-0 silk suture, leaving one end of the suture long for later use. Divide the bile duct by cutting between the 5-0 and 6-0 ligations.
12. Mark the top side of the IHVC and PV with a surgical dye pen to help align the vessels during anastomosis, then clamp the portal vein with a microvessel clamp as distal to the liver as possible.
13. Insert a 20 mL syringe with a 26 G needle into the PV proximal to the micro clamp and perfuse the liver with 10-15 mL of ice-cold heparinized saline; simultaneously, divide the IHVC using spring scissors as close to the right renal vein as possible.
14. Excise the liver using spring scissors by dissecting the PV proximal to the micro clamp, tightening the 5-0 suture previously placed around the SHVC, and dissecting the diaphragm to transect the intrathoracic vena cava.
15. Continue by dissecting the remaining ligaments at the back of the liver and divide the previously ligated adrenal vein. Place the excised liver in cold saline on ice.

3. Auxiliary liver recipient transplant procedure

1. Anesthetize the recipient rat with 5% isoflurane inhalation in an induction chamber. Record the weight of the rat and shave the abdomen with an electric clipper.
2. Position the rat in a supine position on a heated surgery pad with its nose in an anesthesia cone and immobilize the limbs with tape. Apply an eye lubricant, disinfect the abdomen with 75% alcohol, and lower the isoflurane to 2%.
3. Using scissors, make a longitudinal midline skin and muscle incision from the pubis to the xiphoid, then install retractors on both sides of the abdominal wall.
4. Using moist cotton swabs, retract the intestines to the right side of the abdomen and cover them with moistened gauze. Apply another moist gauze to cover the stomach, spleen, and liver, exposing the left kidney and renal vessels.
5. Using a 27 G hydro-dissection needle and blunt tip forceps, separate the left renal vein from the renal artery, carefully removing fat and connective tissue from both vessels.
6. Isolate the gonadal and adrenal veins, and use 6-0 silk to ligate them proximal to the renal vein temporarily. Cauterize, using bipolar forceps, all micro side branches isolating the renal vein and artery between the aorta/VC and the kidney.
7. Mobilize and ligate the ureter with a 6-0 silk suture at the inferior pole. Mark the renal vein and artery with a surgical dye pen to help orientate the vessels during anastomosis and ensure there are no twists.
8. Clamp the renal artery and the renal vein with a microvessel clamp as close to the aorta and VC as possible. Transect the renal artery with spring scissors just past the vessel bifurcation and divide the renal vein about halfway between the VC and kidney. Mobilize the left kidney from the surrounding connective tissue and remove it.
9. Flush both vessels with heparinized saline using a 27 G hydro-dissection needle to remove all remaining blood.
10. With spring scissors, cut a small fish mouth opening in the fork of the renal artery bifurcation to make a funnel-shaped opening and insert an 8 mm stent cut from a 26 G catheter (Figure 2B). Secure the stent with 6-0 silk suture, leaving one end of the suture long for later use.
11. Introduce the donor liver and position it with the PV, IHVC, and bile duct facing the recipient's left renal vein and artery. Using a 9-0 nylon suture, install two stay sutures on opposite sides of the IHVC-renal vein connection.
12. Compare the width of the vessels and make a small fish mouth incision using spring scissors into the face of the renal vein until it has a similar width to the donor's IHVC (Figure 2C).
13. Using a 9-0 nylon suture, anastomose the liver IHVC end-to-end to the renal vein with 9 or 10 running sutures across both front and back walls of the vessel. Alternatively, use a cuff method to complete this anastomosis^3,4.
14. Confirm that placement of the renal artery is beneath the IHVC (Figure 2D), and insert the stent previously placed in the renal artery into the liver portal vein and secure with a 6-0 silk suture, leaving one end of the suture long to attach to the opposite thread on the artery. Draw the ends together to hold each from sliding off the stent.
15. Remove the micro clamp on the renal vein first, then remove the micro clamp on the renal artery.
16. During reperfusion of the liver, use gauze and cotton swabs to apply light pressure around the anastomosis region until a patent anastomosis is achieved (Figure 2D). Remove the temporary ligations previously placed on the adrenal and gonadal veins.
17. Carefully mobilize approximately 10 mm down the end of the left ureter from surrounding connective tissue, leaving a significant amount of fat tissue attached. With spring scissors, make a small incision in the wall of the ureter proximal to the 5-0 ligation previously placed.
18. Insert the polyimide stent previously attached to the bile duct into the small incision made in the ureter wall. Secure with a 6-0 silk suture and tie one end together with the long thread on the bile duct side of the stent, drawing both ends firmly together.
19. Return the intestines back to their natural position (Figure 2E), irrigate with 2-3 mL of saline, and close the abdomen in two layers using 3-0 silk running sutures.
20. Inject 0.1 mg/kg buprenorphine subcutaneously, place the recipient in a clean, heated cage, and monitor recovery for 1-2 h before returning the animal to the animal housing facility.

4. Post-surgical follow-up

1. Starting on day 2 post-surgery, inject allogeneic transplant recipients with heparin (1 IU/g) subcutaneously daily.
2. Starting on day 2 post-surgery, inject syngeneic transplant recipients with heparin (1 IU/2 g) subcutaneously every other day.

Results

Presently, 29 pairs of rats have been used to establish the HALT-PVA protocol, 17 syngeneic transplants, and 12 allogeneic transplants. The syngeneic transplanted livers survived to their designated 8 or 30-day experimental endpoint with a 70% success rate, while allogeneic transplanted livers survived to their designated 3 or 8-day endpoints with a 50% success rate. Failures include rats that died due to surgical complications and auxiliary livers that failed even when the recipient survived.

Discussion

Liver transplantation is the only treatment option for patients with end-stage liver disease, with almost 9,000 LTxs performed yearly in the US¹³. Unfortunately, immunological rejection is seen in up to 25% of LTx recipients, and this rejection is detrimental to the transplanted organ and patient^14,15. To improve outcomes after LTx, the development of innovative models to study organ rejection and implement strategies to decrease rejection...

Materials

Name	Company	Catalog Number	Comments
3-0 Silk Suture	Ethicon	C013D
5-0 Silk ties	Fine Science Tools	18020-50
6-0 Silk ties	Fine Science Tools	18020-60
7-0 Silk ties	Teleflex	103-s
9-0 Polyamide Suture	AROSurgical	T05A09N10-13	Black
Bipolar Cautery	Codman & Shurtleff Inc.	P.H. 234
Buprenorphine HCL	Hospira	409201232
Forceps, Adson-Brown	Fine Science Tools	11627-12	12.5 cm
Forceps, Angled Dumont	Fine Science Tools	11253-25	Medical #5/45 11 cm
Forceps, Suture Tying	Fine Science Tools	18025-10	10 cm
Heparin Sodium Injection, USB	Fresenius Kabi	504015	10,000 USP units per 10 mL
Hydrodissection Cannula	Ambler Surgical	1021E	27 G
Isoflurane	Dechra Vet. Products	17033-091-25
I.V. Catheter	Kendall	2619PUR	26 G x 3/4"
Magnetic Retraction System	Fine Science Tools	18200-50
Micro Clamps	Fine Science Tools	18055-05	6 mm
Micro Clamps	Fine Science Tools	18055-06	4 mm
Micro Clamp Applicator	Fine Science Tools	18057-14	14 cm
Micro Needle Holder	S&T	C-14	14 cm
Microscope	Zeiss	Universal S3	Dual head
Ophthalmic Ointment	Puralube	14590500
Polyimidi Tubing	Cole Parmer	95820-04	OD 0.0215", ID 0.0195", wall 0.0010"
Saline	Baxter	281324	0.9% Sodium Chloride
Surgical Spring Scissors	S&T	SDC-15	Blunt 14 cm
Surgical Spring Scissors	Fine Science Tools	15021-15	Vannas 14 cm