TECHNIQUE OF KIDNEY TRANSPLANTATION IN MICE WITH ANTI-REFLUX URINARY RECONSTRUCTION
( Download pdf )

PAULO N. MARTINS

Department of Surgery, Division of Transplant Surgery, Brigham and Women’s Hospital, Harvard Medical School, Boston, MA, USA

ABSTRACT

     Experimental models of organ transplantation play a crucial role in establishing the principles of transplantation immunobiology. Murine transplant models of vascularized organs are particularly useful for immunobiological studies because there are more immunological tools available. However, the technique of kidney transplant in mice is very challenging. A difficult aspect of this model is urinary reconstruction, which is frequently associated to complications. In this article, the technique of mouse kidney transplantation using an anti-reflux system (modified extravesical ureteroneocystostomy) is described and illustrated for the first time. Although technically demanding, this procedure is feasible and may reduce the incidence of urine leakage and reflux.

Key words: kidney transplantation; experimental model; mouse; microsurgery
Int Braz J Urol. 2006; 32: 713-20

INTRODUCTION

     Transplant models of vascularized organs in the mouse are particularly useful for immunobiological studies because there are more immunological tools available (inbred, transgenic and knockout animals; genetic mapping, monoclonal antibodies, reagents, etc.) (1,2).
     The technique of mouse kidney transplantation was first established by Skoskiewicz & Russel already in 1973 (3), then improved by Kalina (4) and Zhang (5). However, it is currently performed only in a few transplantation centers in the world, due to the anastomoses complexity (renal artery diameter 0.3 mm).

SURGICAL TECHNIQUE

     Male inbred C57BL/6 mice (Jackson Laboratories) weighting 20-30 g were used for kidney transplantations. Animals were housed and cared for in accordance with our institution’s guidelines for experimental animals. In a small series of 20 animals, consecutively operated using this technique, the failure rate was 50% and apparently related to vascular complications.
     Mice were anesthetized with a solution of ketamine (80 mg/kg i.p.) and xylazine (5 mg/kg i.p.) mixed in a normal saline solution. All procedures were performed at 4-25x magnification using a microscope (SMZ800, Nikon, Japan) and standard microsurgical instruments.

Donor Operation
     After induction of anesthesia, the mouse was shaved, fixed on the operating board and the abdomen was prepped with betadine. To assure normal volemia, a saline solution was instilled in the abdominal cavity whenever it became dry. A midline incision from the xyphoid appendix to pubis was performed and exposure was achieved using small retractors to keep liver and bowels away from the left kidney.
     The renal vein and artery were dissected apart under X20 magnification avoiding direct manipulation of the vessels. Left adrenal and testicular vessels were ligated with 10-0 polypropylene sutures and divided. Then, the kidney was separated from the perinephric fat and adrenal gland, the ureter was dissected freely down to the bladder without stripping its fat and cut with a small (1-2 mm) bladder patch. After clamping the aorta above the renal artery, a 30-gauge needle was introduced into the aorta, the inferior (caudal) vena cava was cut and the graft perfused with 1.0 mL cold heparinized saline solution (100U/mL). Renal vessels were cut with a Carrel patch of the aorta and vena cava and the graft was removed and stored in a normal saline solution at 4°C for 20 min. until the time of transplantation (Figure-1).



Recipient Operation
     To increase the success rate, a two-stage procedure was performed, i.e. the left native kidney was removed at the time of grafting and the right native kidney was left in place until day 7, when the right nephrectomy was performed.
     A midline incision from the xyphoid appendix to pubis was performed, and exposure was achieved using small retractors to keep liver and bowels away from the kidney. The renal pedicle was ligated with 7-0 silk suture and the recipient left kidney was removed leaving place for the donor kidney.
     After ligating some lumbar branches, the infrarenal aorta and inferior vena cava were isolated and two loops of 7-0 silk were placed proximally and distally around them and tied to promote homeostasis. Elliptical longitudinal aortotomy and cavotomy were performed between these ties and the vessels were flushed to clear blood inside them. Vascular anastomoses were performed end-to-side to the abdominal aorta and vena cava using a running 11-0 nylon suture (Ethilon, Ethicon, USA). To anastomose the renal artery, two stay sutures (11-0 nylon suture) were placed at the artery’s opposite ends (inferiorly and superiorly). The back wall was sewn first, through a transluminal approach; then the anterior wall, both with a continuous suture. In general, only four to five stitches on each side are necessary to avoid bleeding.
     The vein was anastomosed in the same fashion as the artery. Before suturing the vein, it is important to confirm it is not twisted. Depending on the size of the caval patch, 5 to 7 times the vein was stitched on each side, avoiding pinching the opposite wall. After both anastomoses were performed, the first distal loop was loosened and then the proximal loop. Blood flowed back into the kidney and bleeding was minimal. Saline was usually given subcutaneously proportional to the bleeding (Figure-2).



     Urinary reconstruction was established by suturing the small bladder patch to a cystotomy located on the bladder dome (similar to the Lich-Gregoir technique for extravesical ureteroneocystostomy). The animal should be well hydrated to keep the bladder filled. First, the serosa and muscular layers of the posterolateral aspect of the bladder were divided (5 mm) with the micro-scissors avoiding major vessels and taking care not to enter the bladder. This develops a plane between the muscularis and mucosa allowing the mucosa to bulge out. Then, the bladder mucosa was divided (2 mm) and sutured to the mucosa of the patch from the donor’s bladder with a running 11-0 nylon suture (Ethilon, Ethicon, USA). Then, one interrupted seromuscular suture was placed in each quadrant of the patch, and finally three interrupted 11-0 nylon seromuscular sutures were loosely placed over the patch to close the bladder muscle over the bladder patch and terminal ureter, burying the anastomosis, sealing the first suture and creating a muscular tunnel to reduce the risk of urine reflux (Figure-3).



     Abdominal wall and skin were closed with a continuous 5-0 absorbable suture (Vycryl, Ethicon, USA). A saline 0.5 mL solution was given subcutaneously after the transplant and no antibiotics or heparin were given. Mice were kept under a heating lamp until they were awake.

COMMENTS

     Mouse kidney transplantation is a very demanding microsurgical procedure full of details, which requires long training and meticulous technique. The murine kidney is extremely sensitive to ischemia/reperfusion injury, and to perform a successful operation the time of anastomosis (warm ischemia) should be less than 35 minutes. Most experiments show that in experienced hands the survival rate is between 40 and 70% (3,4). The learning curve is slow because the risk of renal artery thrombosis is very high and difficult to avoid, and due to urological complications.
     In rat kidney transplantation, continuity of urine flow is reestablished most commonly by end-to-end ureter-ureter anastomosis (splinted or non-splinted). On the other hand, kidney transplantation in mice, probably because of the very small diameter of the ureter, ureter-to-ureter anastomosis has not been described. In the literature, urinary reconstruction in mouse kidney transplantation has been performed either by direct implantation of the ureter in the bladder without anastomosis or most commonly by bladder-to-bladder anastomosis using a large bladder patch. The blood supply of the terminal ureter and bladder patch is exclusive from the graft vasculature and prone to ischemia. Not uncommon, the use of a large bladder patch is associated with either bladder necrosis and urine leakage or neurogenic bladder with calculi formation due to bladder denervation (6).
     The most preferred technique for urinary reconstruction in clinical kidney transplantation is extravesical ureteroneocystostomy. Differently from the technique of extravesical ureteroneocystostomy used in the clinic, the author opted for using a small bladder patch since spatulation of the mouse ureter would be difficult. No splint was used since it has been associated with more urological complications (6), and the bladder patch allows enough area to place sutures. The technique described here avoids continuous (running) suture of muscular layer that increases ischemia and the risk of patch necrosis and urine leakage. Water-tightness is ensured by minimal vesical opening and continuous suture. The incidence of vesical calculi depends also on the suture material used for urinary reconstruction (6), and therefore, newer absorbable microsutures are preferred.
     An anti-reflux system, although time-consuming and difficult to perform, may reduce the incidence of urosepsis related to urinary fistulae and reduce the risks of hydronephrosis and chronic pyelonephritis in the long-term. To the author’s knowledge this is the first report on urinary reconstruction in the mouse using an anti-reflux technique. The potential advantages of this technique applied in the murine model still need to be confirmed in the long-term in case-control studies.

ACKNOWLEDGEMENT

     This work was performed at the Charité-University Medicine Berlin, Virchow Clinic, Department of General Surgery and Transplantation, Berlin, Germany.

CONFLICT OF INTEREST

     None declared.

REFERENCES

____________________
Accepted after revision:
August 9, 2006

_______________________
Correspondence address:
Dr. Paulo Ney Aguiar Martins
Division of Transplant Surgery
Brigham and Women’s Hospital
221 Longwood Ave, Rm 309
Boston, Massachusetts, 02115, USA
Fax: + 1 617 732-5724
E-mail: pmartins@partners.org