Orthotopic Rat Kidney Transplantation: A Novel and Simplified Surgical Approach

Summary

The purpose of this manuscript and protocol is to explain and demonstrate in detail the surgical procedure of orthotopic kidney transplantation in rats. This method is simplified to achieve the correct perfusion of the donor kidney and shorten the reperfusion time by using the venous and ureteral cuff anastomosis technique.

Abstract

Kidney transplantation offers increased survival rates and improved quality of life for patients with end-stage renal disease, as compared to any type of renal replacement therapy. Over the past few decades, the rat kidney transplantation model has been used to study the immunological phenomena of rejection and tolerance. This model has become an indispensable tool to test new immunomodulatory pharmaceuticals and regimens prior to proceeding with expensive preclinical large animal studies.

This protocol provides a detailed overview of how to reliably perform orthotopic kidney transplantation in rats. This protocol includes three distinctive steps that increase the probability of success: perfusion of the donor kidney by flushing through the portal vein and the use of a cuff system to anastomose the renal veins and ureters, thereby decreasing cold and warm ischemia times. Using this technique, we have achieved survival rates beyond 6 months with normal serum creatinine in animals with syngeneic or tolerant kidney transplants. Depending on the aim of the study, this model can be modified by pre- or posttransplant treatments to study the acute, chronic, cellular, or antibody-mediated rejection. It is a reproducible, reliable, and cost-effective animal model to study different aspects of kidney transplantation.

Introduction

Historically, the first transplant rejection studies were performed by Brent and Medawar using skin transplants in rodents¹. It soon became clear that skin has distinct immunological features, making it a highly immunogenic organ that is different in rejection from other vascularized solid organs². Rat studies of solid organ transplant rejection are habitually limited to heart, liver, and kidney transplants. Although each of these organs is suitable to study rejection, there are advantages and disadvantages to each of them. Heart transplants are often transplanted into the abdomen and anastomosed to the aorta and vena cava, with the recipient’s native heart in place³. This does not recreate human clinical, anatomical, and physiological conditions. Additionally, hearts are very sensitive to cold ischemia and have to be reperfused preferentially within 1 h in order to be able to recover their function⁴. Liver transplants are generally considered to be surgically more challenging and time-sensitive to perform. After removing the native liver, the donor liver has to be implanted and reperfused within 30 min as the recipients cannot last longer without a functioning liver⁵. The hepatic artery, portal vein, and especially the bile duct reconstruction requires refined surgical skills. Besides the surgical challenges, the liver is known to possess tolerogenic properties and rodents and humans can become operationally tolerant^6,7,8. The kidney, unlike the aforementioned organs, can be transplanted in an orthotopic fashion, is known to be an immunogenic organ with consistent, reproducible rejection episodes (if not immunosuppressed), and allows for prolonged cold ischemia times of several hours. This makes the rat kidney transplant an ideal model for studying allograft rejection and tolerance.

Kidney transplantation (KT) is the preferred choice of treatment for patients with end-stage renal disease. Over the last few decades, short-term survival outcomes after KT have improved dramatically, but long-term survival outcomes are stagnant⁹. Conventional immunosuppressive regimens remain the standard anti-rejection therapy. However, the chronic use of immunosuppressive therapies causes significant morbidity and mortality, such as nephrotoxicity, diabetes, and secondary malignancies^10,11,12. In the long-term, chronic antibody- and cellular-mediated rejection threaten graft survival, with limited therapeutic options available.

A major goal in transplantation is the induction of transplant tolerance in order to obviate the need for chronic immunosuppression. The rat KT model is a robust tool to investigate the immunological rejection process and to evaluate new approaches to immunomodulation and transplant tolerance. The rat also serves as a suitable model to study acute and chronic cell- and antibody-mediated rejection^{13,14,15,16,17}. This surgical model has proven to be a reliable, reproducible, and cost-effective tool to study various aspects of allograft rejection and tolerance. It is often used to test novel tolerance-inducing protocols prior to undertaking expensive and cumbersome large-animal studies. Performing KT in rats requires extensive surgical training and expertise to reach survival rates of >90%. In this manuscript and in the accompanying instructional video, we provide a step-by-step outline for orthotopic KT in the rat, as successfully performed for many years at our institution.

Prior to starting any procedure, donor and recipient selection is critical and depends on the nature of the experiment. Ideally, donors and recipients should weigh between 220–260 g and be between 8–12 weeks of age. Animals under 220 g have small-diameter arteries, veins, and ureters, making the anastomosis in the recipient particularly challenging. Minor blood loss can cause hypovolemia and lead to death in smaller animals. Animals heavier than 260 g display more fat around their vessels, and vessel isolation will require more operative time and increase the cold ischemia time.

Protocol

Lewis (RT1¹) and Dark Agouti (DA) (RT1A^a) rats were purchased from commercial vendors (see the Table of Materials). These fully MHC-mismatched strains are often used to study acute renal allograft rejection. All animals were housed and maintained according to the National Institutes of Health’s (NIH) guidelines in a specific pathogen-free facility at the Johns Hopkins University. All procedures were approved by the institutional animal care and use committee.

1. Donor procedure

1. Prepare and autoclave all surgical instruments to be used in this procedure as a means of sterilization and use disposable sterile gloves to prevent infectious complications.
2. Anesthetize the donor rat by isoflurane inhalation (induction at 3%–4% and maintenance at 1%–2%) for the rest of the procedure. Give all donor and recipient animals preemptive buprenorphine subcutaneously at 0.1 mg/kg body weight for analgesia.
3. Now, place the rat in a supine position and immobilize the limbs with sterile masking tape.
4. Use a mechanical clipper to remove hair from the abdominal area.
   1. Apply an eye lubricant and use sterile gauze soaked in povidone-iodine, followed by gauze soaked in isopropyl alcohol, to sterilize the surgical field.
   2. Prior to the first incision, make sure that the rat is adequately anesthetized by checking the absence of the toe pinch withdrawal reflex.
5. Using scissors, start off by making a large longitudinal midline skin and muscle incision from the symphysis pubis to the xiphoid, and enter the peritoneal cavity.
6. Insert two retractors on either side of the abdominal wall in order to expose the intra-abdominal cavity.
7. Cover the intestine with a moist sterile gauze and shift it to the right lateral side of the abdomen, exposing the aorta, vena cava, and left kidney. Apply 1 mL of preheated saline with a 1 cc syringe to keep the intestines and the abdominal organs moist and at a normal temperature.
   1. Apply a second moist gauze to cover and mobilize the stomach and spleen cranial to the kidney and a small moist gauze to cover the exposed kidney (Figure 1A).
8. Use microsurgical dissecting forceps to isolate and mobilize the left renal artery and vein from the connective tissue and each other. Isolate the left renal vein by cauterizing the left gonadal vein and isolate the left renal artery by cauterizing the adrenal artery. After that, mobilize the aorta and vena cava superior and inferior of the left renal pedicle by dissecting the connective tissue with dissecting forceps (Figure 1B).
9. Divide and mobilize the ureter from the connective tissue using dissecting forceps, and make a diagonal incision at a length of 2 cm measured from the renal pelvis, using microscissors. Insert a polyamide cuff (see Table of Materials) halfway into the ureter and secure the cuff by placing a knot with 8-0 silk suture (Figure 1C).
   ​NOTE: It is important not to remove all fat and connective tissue from the ureter, as they provide protection against obstruction caused by adhesions, and their removal may cause ureteral necrosis. Pay extra attention to preserving the vessel supplying oxygen to the ureter.
10. Mobilize the left kidney by separating it from the perinephric fat using dissecting forceps or microscissors. Leave the adipose capsule of the kidney attached and use that site for handling the kidney.
    1. Expose the inferior vena cava.
11. Administer 200 units of heparin using a syringe with a 27 G needle through the penile vein. Pressure the site of injection with a cotton swab for at least 1 min to prevent bleeding.
12. Identify the portal vein (pv) and inferior vena cava (ivc) (Figure 1D). Flush the kidney by injecting 50 mL of cold saline mixed with 500 units of heparin into the portal vein using a 16 G needle (Figure 1E). Before flushing, cut the inferior vena cava at the infrahepatic level and the portal vein caudal at the needle insertion site to allow the blood to exit the circulation. Start flushing the kidney by gradually infusing the saline solution. Observe a change of color of the kidney from dark red to a uniform grey and pale color (Figure 1F).
13. After flushing, ligate the renal artery and vein proximal to the aorta and vena cava and place the flushed kidney in a Petri dish in cold saline on ice. Figure 2A represents the schematic overview of the donor procedure.
14. Once the kidney is in cold saline, fix and immobilize the handle of the vein cuff (see the Table of Materials) and gently pull the renal vein through the cuff. Then, fix the renal vein over the cuff by placing three knots using eight-zero silk suture (Figure 2B).                                                                       
    NOTE: Pay special attention to the orientation of the vein while securing it in place. Rotated veins cause an obstruction of the blood flow and lead to thrombosis.

2. Recipient procedure

1. Repeat steps 1.1–1.11 from the donor procedure.
2. Place two atraumatic micro-vessel clamps on the left renal artery and vein proximal to the aorta and vena cava (Figure 3A).
3. Ligate the recipient renal vein proximal to the inlet of the kidney. Flush the renal vein with heparinized saline to remove all the remaining blood out of the vessel.
4. Slide the ligated renal vein over the cuffed renal vein previously positioned in the donor kidney and secure it with an 8-0 silk suture (Figure 3B). Maintain the same positional orientation when securing the renal vein over the cuff.
5. Ligate the ureter at the level of the lower pole of the left kidney. Mobilize the kidney from the perinephric fat.
6. Ligate the renal artery proximal to the inlet of the recipient kidney. Flush it with heparinized saline to remove any excess blood in the vessel. Perform an end-to-end anastomosis of the renal artery with 8 to 10 interrupted sutures using a 10-0 nylon suture (Figure 3C). Maneuver the artery by using the adventitial layer.
7. Remove the vessel clamps to reinitiate the reperfusion of the kidney. Start by removing the clamp on the vein followed by the clamp on the artery (Figure 3D). Use a sterile cotton swab to lightly pressure any oozing areas around the anastomosis region. A few minutes should be sufficient to achieve a patent anastomosis.
8. Briefly observe the kidney to assess for adequate perfusion. Immediately after reperfusion, the kidney should change color and gradually regain its natural dark red color after a few minutes (Figure 3E). Visible peristalsis of the ureter and on-site urine production are sometimes observed.
9. Finish by inserting the exposed tip of the ureteral cuff into the recipient ureter and secure the recipient ureter with an 8-0 silk suture (Figure 2C and Figure 3F).
10. In order to keep the donor and recipient ureters in position, tie off the ends of each ureter side to each other.
11. Optionally, the right kidney can be nephrectomized by tying off the right renal artery and vein with a 4-0 silk suture and removing the kidney.
12. Remove all gauzes from the abdominal cavity, return all the organs to their natural position, squirt 1 mL of saline over the intestines to keep them moist, and close the abdomen by using a 4-0 absorbable suture on the rectus muscle and a 4-0 silk suture to close the skin layer in an interrupted fashion.

3. Postoperative care

1. Place the animal in a clean cage with access to ad libitum water and food and allow for recovery on a 37 °C heating pad.
2. Inject 0.1 mg/kg buprenorphine subcutaneously for analgesia and monitor the animal for recovery. Additional analgesics administration may be needed in the next few days, depending on signs of discomfort or pain. Antibiotics are not routinely administered, as infectious complications are rare.
3. Observe the recovery for 1–2 h before returning the animal back to the animal facility. Inspect the animal 2x–3x a day for the first 24 h, followed by a daily inspection. Pay attention to signs of pain and distress, oral intake, and urinary output.
4. Remove the stitches 7–10 days after the operation.

Results

We performed syngeneic (N = 5) and allogeneic kidney transplants (N = 5). Animals with a syngeneic transplant achieved long-term survival without any immunosuppressive treatment. Animals that received an allogeneic transplant without immunosuppression rejected their graft and succumbed to renal failure with a median survival of 8 days (Figure 4A). Mean serum creatinine increased modestly in the syngeneic group while it increased by 14-fold in the allogeneic...

Discussion

In this manuscript, we describe the surgical method for orthotopic KT in rats in detail, including all the necessary equipment needed to perform this procedure (Figure 5). In 1965, Fisher and Lee published the first report on KT in rats, which became the start of an exciting investigative field¹⁸. Since then, many modifications have been introduced to improve the reproducibility of this model. It has served as an effective animal model for studying ischemia-reperfusio...

Materials

Name	Company	Catalog Number	Comments
Buprenorphine HCL	Reckitt Benckiser Healthcare UK	NDC12496-0757-5
Dissecting forceps, curved	Zhenbang, China		11cm Flat handle
Heparin sodium injection USP	Sagent Pharmaceuticals	NDC25021-400-10
Micro-forceps, straight, smooth	Jingzhong, China	WA3010
Micro needle holder	Jingzhong, China	WA2010
Micro vessel clamps	Jingzhong, China	WA40120
Micro spring sciccor 1	ROBOZ	RS-5620
Micro spring sciccor 2	F.S.T.	91501-09
Micro spring sciccor 3	Zhenbang, China		8.5cm Vannas,curved
Prograf (Tacrolimus/FK506)	Astellas
Rats	Charles River & Taconic Biosciences	LEW/Crl & DA-M
Shaver	Wahl	79600-2101
Suture 4-0	Ethicon	J304H
Suture, 4-0	Ethicon	683G
Suture, 10-0	Ethicon	2820G
Syringes & Needles	BD
Thread, 8-0	Ashaway	75290
Ureteral cuff	Microlumen	160-1	Polymide Tubing, Diameter 0.41 mm
Venous cuff	Intramedic BD	7441	PE-200 Non-radiopaque polyethylene tubing ID: 1.4 mm, OD: 1.9 mm