A Modified Surgical Technique for Kidney Transplantation in Mice

Summary

This protocol presents a new surgical technique of mouse kidney transplantation focusing on a modified arterial anastomosis strategy. A vascular suture technique including a simple and safer ureter-bladder anastomosis method is also presented. These modifications shorten the operation time and improve the success rate of the mouse kidney transplantation procedure.

Abstract

Kidney transplantation in mice is a complicated and challenging surgery procedure. There are very few publications demonstrating the key steps of this operation. Therefore, this article introduces the technique and points out the surgical caveats associated with this operation. In addition, important modifications in comparison to the conventional procedure are demonstrated. Firstly, a patch of the abdominal aorta is cut and prepared so that the proximal bifurcations of the renal artery, including the ureteral artery are transected together with the donor kidney en bloc. This reduces the risk of a ureter necrosis and avoids the development of a urinary tract occlusion. Secondly, a new method of the vascular anastomosis is demonstrated that allows the operator to flexibly increase or decrease the size of the anastomosis after renal transplant reperfusion has already been initiated. This avoids the development of vessel strictures and intraabdominal bleeding. Thirdly, a technique that enables the anastomosis of the delicate donor ureter and the recipient bladder that does not cause a trauma is shown. Adopting this protocol can shorten the operation time and reduces the damage to the recipient's bladder, thereby significantly increasing the operation success rate for the recipient mice.

Introduction

Since Sakowitz et al. developed mouse models of kidney transplantation in 1973 for the first time¹, it has proven as an important experimental tool to study the mechanisms of transplant ischemic injury and alloimmune rejection as well as for developing new treatments aimed to prolong allograft survival and possibly to achieve immunological tolerance. However, the surgical technique has proven to be complex and very demanding, sometimes having complications such as vascular anastomotic strictures leading to prerenal non-immunological kidney transplant failure², postrenal failure caused by ischemia and subsequent necrosis of the transplanted ureter, strictures of the anastomosis of the transplanted ureter and/or the recipient's urine bladder leading to a disruption of the urinary outflow. All of these are reasons why renal transplantation in mice has not been further developed and is therefore not widely used. Establishing an effective and long-term stable mouse kidney transplantation model without vascular and urinary tract complications still has irreplaceable significance for many studies in the transplant field with focus on the renal immune mediated but also infectious diseases³. In addition, compared with other organ transplants in murine models such as lung, heart, and intestinal transplantation^4,5, the mouse kidney transplantation model offers a chance for studying long-term survival even in the setting of major histocompatibility antigen disparity^3,6. It has also been shown that in the same setting of donor-recipient strain combinations different organ transplants such as heart or kidney are characterized by different dynamics and onsets of allograft rejection³. Furthermore, from the nephrological point of view, it is a more suitable model for studying parenchymal mediated immune regulatory mechanisms in the context of acute and chronic rejection events than simple skin transplant experiments.

On the basis of previous reports on the surgical technique of kidney transplantation in mice^3,7,8,9, we here demonstrate the following reliable improvements that have been successfully applied during the past 10 years within our group^10,11,12: Firstly, the ureteral artery is safely conserved as the renal artery is resected en bloc together with the respective part of the abdominal aorta. Second, a new, simple, and rapid technique of a knotless vascular anastomosis in which the final stitch of anastomosis is not tied with the end of the upper tie like the traditional approach but remains free. This technique enables to increase or decrease the size of the anastomosis after renal reperfusion to avoid vessel stricture and intraabdominal bleeding. Third, 21 G and 30 G syringe needles were used as an auxiliary puncture guiding tool in order to implant the donor ureter into the recipient's bladder wall reducing the damage to the recipient's bladder and facilitating the formation of stricture free anastomosis.

In this report, we also compared the traditional, widely used technique with the modified one that is established in our laboratory and found no significant difference in the degree of renal tubular atrophy and kidney transplant interstitial tissue fibrosis. In previous studies, we additionally compared the results of this new technique with the conventional method in terms of local bleeding, thrombosis, time for performing the vessel anastomosis and survival rate. We found improvements such as significant reductions of local thrombosis events (1.1% versus 6.6%), a reduced time for the anastomosis procedure, and a highly reproducible kidney syngeneic graft long-term survival (95% versus 84% with the classical approach)¹⁰.

Protocol

All animal experiments were conducted according to the guidelines from the directive 2010/63/EU of the European Parliament on protection of animals used for scientific purposes (Animal ethics card: Lower Saxony Ministry of Food and Drug Safety, #33.9-42502-04-11/0492). Conduct procedures using sterile surgical instruments and consumables (autoclaved) and try to keep the operating area as sterile as possible.

NOTE: C57BL/6J male mice served as donors and recipients (syngeneic transplant model) while Balb/c mice served as kidney allograft recipients (model for studying acute allograft rejection model⁹). Mice were aged between 8-12 weeks, weighed ~25-30 g at transplantation and were housed under standard conditions. Data reported in this manuscript were generated by four surgeons experienced in mice surgery.

1. Preparatory steps

1. For surgery, use a set of microscopic instruments, including a micro-scissor, micro-forceps, a needle holder, micro hemostatic clamps, and an electrosurgical pen. Perform sutures using 7/0er, 10/0er, or 4/0er nylon monofilament.
2. For anesthesia, place the mouse into the box for inhalation of isoflurane (2%) for about 40-60 s in order to induce unconsciousness.
3. Once the mouse is anesthetized, weigh the mouse.
4. According to the mouse´s weight, apply an intraperitoneal injection of ketamine (100 mg/kg) + xylazine (10 mg/kg) + acepromazine (2 mg/kg) to anesthetize the mouse¹³. Confirm that the mouse is anesthetized by observing a lack of response to a toe pinch.
5. When anesthesia has taken effect, clip the abdominal fur. Then, fix the mouse on the operation table by loosely immobilizing the limbs with a sterile masking tape.
6. Disinfect the mouse's abdomen after placing the mouse on the operation table. Perform disinfection using alternating scrub of povidone iodide (iodophor) and alcohol, three times (use concentric pattern, start scrub in the middle of the abdomen and move outward), and then properly drape the mouse using a fenestrated surgical towel.
7. Apply eye ointment and maintain sterility throughout the procedure.
   ​NOTE: Antibiotics are not recommended throughout the procedure as these substances may influence immunological responses.

2. Donor operation procedure

1. Use scissors to cut the skin and perform a cross abdominal incision of about 3-4 cm. Cut the muscles of the abdominal wall. Cover and cautiously move away the viscera with a saline imbibed gauze.
2. Use a cotton swab to gently remove the intestines, stomach, and spleen toward the right side (from point of view of the mouse), cover and cautiously move away the viscera with a saline imbibed gauze.
3. Use micro forceps to expose the left kidney, aorta, and inferior vena cava (IVC).
4. Use an electrosurgical pencil to cauterize the left lumbar veins, including their underlying branches and other small vessels along with the left adrenal vessel, carefully.
5. Use micro scissors and forceps to dissect the left ureter and cautiously mobilize it from the surrounding tissue. Clean cut it close to the urinary bladder. Mobilize the aortic region between the left and right renal arteries approximately 2 mm in length.
6. Use micro forceps to separate the infrarenal inferior vena cava (IVC) and aorta, and then use curved forceps to pass under the aorta to place a loose tie of 7-0 silk suture around this vessel.
7. Cross clamp the area of the aorta below the right renal artery and the inferior vena cava (IVC) using two 5 mm microvascular clamps.
8. Transect the left renal vein from the vena cava.
9. Use a syringe to flush the aorta with 1 mL of heparin saline solution (60 U/mL).
10. Use micro forceps to tighten the ligature applied at step 2.5. Then, cut the aorta below the ligature as well as below the proximal clamp. With this, the proximal bifurcations of the renal artery (please note that the arterial opening must be cut neatly, otherwise it will affect the anastomosis) and the ureteral artery are included and transected en bloc. Prepare carefully, so that the delicate ureteral artery is completely preserved.
11. Use the electrosurgical pencil and forceps to free the left kidney and associated vessels completely by cautiously cauterizing all vessel surrounding tissue. Remove the kidney and store it in saline solution at 4 °C.
12. Euthanize the anesthetized donor mouse by decapitation.

3. Recipient operation procedure

1. Perform the initial surgical steps (including anesthesia and sterilization, see steps 1.1 to 1.7) as described for the donor mouse.
2. Use scissors to open the abdomen via a median incision (about 2.5 cm in length), and then cover the abdominal organs with a wet gauze using saline solution.
3. Carefully preserve the infrarenal aorta and inferior vena cava (IVC) and make sure every large vessel branch is cauterized. Use the electric cautery as well to dissect the left ureter carefully at a position proximal to the kidney pelvis. Then, remove the left kidney.
4. Use micro forceps and cotton buds to expose the abdominal aorta and inferior vena cava and detach them from the surrounding adipose tissue (approximately over 4 mm in length).
5. Use two microvascular clamps and position them proximally and distally on both the inferior vena cava and the abdominal aorta simultaneously.
6. Use a micro needle holder to guide a 10/0 monofilament (made of synthetic fiber with a smooth surface) suture needle, which is placed through the aorta wall in a proximal to distal manner.
7. Achieve an elliptical arteriotomy of approximately 1 mm with a gentle upward traction of the suture, while cutting directly below the lower face of the needle with fine, curved scissors.
8. Use micro scissors to cut the inferior vena cava (IVC) longitudinally with sufficient length of approximately 1.5 mm. Position this incision slightly below its aortic counterpart.
9. Perform the donor and recipient aorta anastomosis in an end-to-side manner. Place the donor kidney on the right side of the recipient's inferior vena cava aligning the cuff of the donor's abdominal aorta with the anastomosis of the recipient's abdominal aorta.
10. Use a micro needle holder and two separate 10-0 sutures to stitch the proximal and distal ends of the anastomosis.
11. After tying, leave the two long sutures, including the needle, in place. Sew the left side of the aortal wall of the anastomosis continuously with two evenly spaced stitches in a distal-proximal direction.
12. After the last stitch, guide the suture through a partial thickness of the vessel wall above the upper stay suture tie.
13. Use micro forceps to simultaneously exert gentle traction to the short end of the lower suture tie.
    NOTE: In this new knotless technique, the last stitch is not tied to the short end of the upper tie.
14. Use micro forceps to turn over the transplanted kidney to its normal position. Now continuously sew the right wall of the aortal anastomosis using three stitches in a proximal to distal manner.
    NOTE: Compared with the conventional surgical technique^7,8 the last suture is merged with the distal tie nearby. Do not tie it to the end of the lower suture, cut it to leave a free length of 2-3 mm instead.
15. Perform the venous anastomosis using the same suturing procedure as previously described with the difference that four to five stitches are needed for each side of the anastomosis. The final stitch is left as a free end of similar length similar to the aortal anastomosis described above.
16. After completing both anastomoses, use a dry swab to exert gentle pressure toward the sutured area for about 10-20 s.
17. Use a clip applicator forceps to remove both clamps, first the lower then the upper. Rinse the abdominal cavity with 0.9% sodium chloride at a temperature of 38.0 °C.
18. Observe the reperfusion of the transplanted kidney.

4. Ureteral implantation

1. Use a micro needle holder to penetrate through the recipient's urine bladder with a 10/0 suture (straight needle) and insert it into a 21 G needle lumen for guidance (see Supplementary Figure 1a).
2. Now guide the 21 G needle to stitch a hole at the place of the previous needle application (Supplementary Figure 1b).
3. Pull out the 21 G needle (Supplementary Figure 1c).
4. Use a micro needle holder and 10/0 suture to stitch (no tie) the trimmed ureter end and perforate the bladder with this 10/0 suture again at the place of its entry (Supplementary Figure 1d).
5. Use a micro needle holder to tow the 10/0 filament and the ureter into the urine bladder through the constructed hole (Supplementary Figure 1e).
6. Use a micro needle holder and another 10/0 suture to anastomose the donor's ureter to the recipient's urine bladder. Here, connect the outer membrane of the ureter to the outer membrane of the bladder wall, and perform intermittent sutures with 3 to 4 stitches. Finally, pull out the traction suture (Supplementary Figure 1f).
7. Use forceps to place the intestines back into the abdominal cavity. Perform two-layer sutures (first the abdominal muscles followed by the skin) to close the abdominal wound using a 4/0 filament.
8. Place the transplanted mice into an oxygen and temperature-controlled chamber for recovering after surgery.
9. For postoperative analgesia, directly give Metamizol 200 mg/kg per os after operation.
   Four and 16 h after operation give Metamizol 200 mg/kg per os plus Carprofen (5mg/kg) s.c. In the further follow up, apply Carprofen (5 mg/kg) s.c. to the transplanted mice every 24 hours for three consecutive days after operation¹³. If there are any signs of an insufficient analgesia buprenorphine 0.05 mg/kg is additionally given every 8 h s.c.

5. Contralateral nephrectomy and sacrifice of the recipient mouse

NOTE: Perform contralateral nephrectomy of the recipient mouse 5 days after transplantation.

1. Perform the contralateral nephrectomy of the transplanted mouse 5 days after transplantation under anesthesia. Ligate and cut the recipient's autologous right renal arteries and veins, remove the right kidney and close the abdominal cavity. The postoperative care and analgesia are the same as described before (see step 4.7).
2. Raise and record the state of the mouse. Provide the transplanted mouse postoperative analgesia, food, and water supply.
3. Four weeks after transplantation, sacrifice half of the transplanted mice and perform H&E staining for their kidney transplants.
4. 12 weeks after transplantation, sacrifice the remaining mice and perform Masson Gold staining of these kidney transplants.

Results

Four weeks after transplantation, both the modified technique as well as the conventional technique displayed moderate signs of renal tubular atrophy^14,15 when compared to the native recipient contralateral kidneys (Figure 1). The degree of the renal tubules atrophy demonstrated no significant difference between the two different techniques. Masson Goldner's trichrome staining^14,15...

Discussion

While the skin transplantation model in mice is simple and easy to perform to study alloimmune rejection events, the surgical techniques for investigating more specifically the alloimmune-related inflammatory alterations after heart¹⁶ and kidney transplantation¹⁰ has been proven to be complex and very demanding. From the transplant nephrologist's point of view, the establishment of an effective and long-term stable mouse renal transplantation model has still an irreplac...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
30G-needles	Braun	456300	-
acepromazine	CP Pharma	Tranquisol P	-
Bepanthen eye ointment	Haus-Apotheke	PZN 01578675	-
Bonn Micro Forceps	FST	11083-07	-
Box for insulation and oxygen supply device	RUSKINN	INVIV	-
C57BL/6J  mice	Charles River. Germany	no catalog number	-
Carprofen	Zoetis	Rimadyl 50 mg/ml	-
CATHETER-FEP 26G	TERUMO	Surflo-W	-
Clip Applicator Forceps Style	FST	18057-14	-
Curved forceps	WPI	14114-G	-
Cutasept skin disinfection	VWR	BODL980365	-
Dehydrator	DIAPATH	Donatello	-
electrosurgical pen	Bovie	CHANGE-A-TIP	-
Embedding machine	Wuhan Junjie Electronics Co., Ltd	JB-P5	-
Ethanol	Sinopharm Group Chemical Reagent Co. LtD	100092683	-
Frozen platform	Wuhan Junjie Electronics Co., Ltd	JB-L5	-
gauze pads, cotton swabs	Lohmann-Rauscher	13353	-
Glass slide	Servicebio	G6004	-
HE dye solution set	Servicebio	G1003	-
Heating mat	THERMO MAT PRO 30W	HTP-30	-
hemostatic sponge	CuraSpon	J1276A	-
heparine-solution	Haus-Apotheke	PZN 03029820	-
ice box	PETZ	No Catalog Number available	-
Imaging system	Nikon	Nikon DS-U3	-
Inhalation anesthesia device	GROPPLER	BKGM 0616	-
isoflurane	CP Pharma	Isofluran CP 1 ml/ml	-
ketamine	Zoetis	no catalog numer	-
Masson dye solution set	Servicebio	G1006	-
metamizole	WDT	no catalog numer	-
Micro scissors	FST	15000-00,15000-10	-
Micro Serrefine ( Clamp ) Angled / 16 mm	FST	18055-06	-
Microscope	Leica	LEICAMZ6	-
Microscope light	SCHOTT	KL2500LED	-
Neutral gum	SCRC	10004160	-
Oven	Tianjin Laibo Rui Instrument Equipment Co., Ltd	GFL-230	-
Pathology slicer	Shanghai Leica Instrument Co., Ltd	RM2016	-
Saline solution (NaCl 0.9 %)	Haus-Apotheke	PZN 06178437	-
scissors	Peha Instruments	991083/4	-
Slides	Servicebio		-
small Petri dish	Sarstedt	8,33,900	-
straight forceps	WPI	14113-G	-
surgical tape	BSN	4120	-
Suture Tying Forceps - 10 cm	FST	18025-10	-
Sutures(10-0)	Medtronic	N2540	-
Sutures(4-0)	ETHILON	V4940H	-
Sutures(7-0)	ETHILON	1647H	-
Syringe (0,3 mL)	BD	324826	-
Syringe (1 mL)	BD	320801	-
Tissue spreader	Zhejiang Kehua Instrument Co., Ltd	KD-P	-
Upright optical microscope	Nikon	Nikon Eclipse E100	-
xylazine	Bayer	Rompun	-
Xylene	Sinopharm Group Chemical Reagent Co. LtD	10023418	-