Orthotopic Rat Kidney Transplantation: A Novel and Simplified Surgical Approach

Our laboratory at Johns Hopkins has been performing small animal kidney and liver transplantation with over two decades of experience. This specific protocol outlines critical steps that are necessary for successfully performing orthotopic kidney transplantation in rats. To increase the probability and reproducibility of success, our protocol includes three distinctive steps, perfusion of the donor kidney through the portal vein and the use of a cuff system to anastomose the renal veins and ureters.

Demonstrating this procedure will be Dr.Le Qi, a graduate student in our laboratory. To begin this procedure, anesthetize the donor rat as described in the text protocol. Place the rat in a supine position and immobilize the limbs with sterile masking tape.

Apply an eye lubricant and sterilize the surgical field as described in the text protocol. Prior to the first incision, make sure that the rat is adequately anesthetized by checking the absence of the toe pinch withdrawal reflex. 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.

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 one milliliter of preheated saline with a one cc syringe to keep the intestines and the abdominal organs moist and at a normal temperature. Apply a second moist gauze to cover and mobilize the stomach and spleen cranial to the kidney.

Then add a small moist gauze to cover the exposed kidney. Use microsurgical dissecting forceps to isolate and mobilize the left renal artery and vein from the connective tissue and each other. Now 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 to the left renal pedicle by dissecting the connective tissue with dissecting forceps. Divide and mobilize the ureter from the connective tissue using dissecting forceps, then use micro scissors to make a diagonal incision at a length of approximately two centimeters measured from the renal pelvis. Insert a polyamide cuff halfway into the ureter and secure the cuff by placing a knot with 8-0 silk suture.

Mobilize the left kidney by separating it from the perinephric fat using dissecting forceps or micro scissors. Leave the adipose capsule of the kidney attached and use that site for handling the kidney. Now administer 200 units of heparin using a syringe with a 27 gauge needle through the penile vein.

Pressure the site of injection with a cotton swab for at least one minute to prevent bleeding. Identify the portal vein and inferior vena cava. Expose the inferior vena cava.

Prepare to flush the kidney by injecting 50 milliliters of cold saline mixed with 500 units of heparin into the portal vein using a 16 gauge needle. Before flushing, cut the inferior vena cava at the infrahepatic level and cut 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 color change of the kidney from dark red to a uniform gray or pale color. After flushing, ligate the renal artery and vein proximal to the aorta and vena cava. Place the flushed kidney in a Petri dish containing cold saline on ice before removing the donor's body from the surgical field.

Once the kidney is in cold saline, fix and immobilize the handle of the vein cuff and gently pull the renal vein through the cuff. Then fix the renal vein over the cuff by placing three knots using 8-0 silk suture. To begin the recipient procedure, repeat the steps from the donor procedure up to heparin administration.

Then place two atraumatic microvessel clamps on the left renal artery and vein proximal to the aorta and vena cava. 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.

Slide the ligated renal vein over the cuffed renal vein previously positioned in the donor kidney. Then secure the vein with an 8-0 silk suture. Maintain the same positional orientation when securing the renal vein over the cuff.

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. Now ligate the ureter at the level of the lower pole of the left kidney.

Mobilize the kidney from the perinephric fat and remove the native kidney. Perform an end-to-end anastomosis of the renal artery with 8-10 interrupted sutures using a 10-0 nylon suture. Maneuver the artery by using the adventitial layer.

Remove the vessel clamps to re-initiate the re-perfusion of the kidney. Start by removing the clamp on the vein followed by the clamp on the artery. Use a sterile cotton swab to add light pressure to any oozing areas around the anastomosis region.

A few minutes should be sufficient to achieve a patent anastomosis. Briefly observe the kidney to assess for adequate perfusion. Immediately after re-perfusion, the kidney should change color and gradually regain its natural dark red color after a few minutes.

Visible peristalsis of the ureter and onsite urine production are sometimes observed. Finish by inserting the exposed tip of the ureteral cuff into the recipient ureter and secure the recipient ureter with an 8-0 silk tie. In order to keep the donor and recipient ureters in position, tie off the ends of each ureter side to each other.

Optionally, nephrectomize the right kidney by tying off the renal pedicle with a 3-0 silk suture. Remove all gauzes from the abdominal cavity and return all the organs to their natural position. Then squirt one milliliter of saline over the intestines to keep them moist.

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. Place the animal in a clean cage with access to ad libitum water and food and allow for recovery on a 37 degree Celsius heating pad. Observe the recovery for one to two hours before returning the animal back to the animal facility.

Animals with a syngeneic kidney transplant achieved long-term survival without any immunosuppressive treatment. Conversely, animals that received allogeneic kidney transplant without immunosuppression rejected their graft and succumbed to renal failure with a median survival of eight days. Mean serum creatinine increased modestly in the syngeneic group while it increased by 14-fold in the allogeneic group.

Upon explantation, the macroscopic view of the syngeneic kidney allograph did not show any abnormalities. The kidney color and internal structures remained intact. In contrast, kidney allographs of rejected animals presented red hemorrhagic patches with the destruction of the internal structures.

Hematoxylin and Eosin stains of syngeneic grafts shown thin glomerular capillary loops with normal numbers of endothelial and mesangial cells. Rejected allographs displayed destroyed glomerular structures with signs of inflammation and tubulitis. CD8 positive staining was performed to confirm T-cell mediated rejection.

While syngeneic allographs showed very few CD8 positive T-cells, the rejected allographs showed a significantly higher number of CD8 positive cells in and around the glomeruli and tubuli confirming T-cell mediated rejection. This model provides significant insight into the immunological phenomena of rejection, tolerance, and donor-host interactions paving the way for the development of therapeutical clinical interventions in solid organ transplantation.