Transplantation and Re-transplantation of Mouse Kidney Grafts to Study Local Immune Responses

Daqiang Zhao Institute of Organ Transplantation, Tongji Hospital, Tongji Medical College, Huazhong University of Science and Technology, Wuhan, Hubei 430030, China. Jiangqiao Zhou Department of Organ Transplantation, Renmin Hospital of Wuhan University, Wuhan, Hubei 430060, China. Adham Abu Ali Department of Surgery, University of Pittsburgh, School of Medicine, Pittsburgh, PA 15261, USA. Roger Tieu Medical Scientist Training Program, University of Pittsburgh, School of Medicine, Pittsburgh, PA 15261, USA. Fadi G. Lakkis Thomas E. Starzl Transplantation Institute, University of Pittsburgh, School of Medicine, Pittsburgh, PA 15261, USA. Martin H. Oberbarnscheidt Thomas E. Starzl Transplantation Institute, University of Pittsburgh, School of Medicine, Pittsburgh, PA 15261, USA. Amit Tevar Department of Surgery, University of Pittsburgh, School of Medicine, Pittsburgh, PA 15261, USA. Zhishui Chen (  zschen@tjh.thumb.edu.cn ) Institute of Organ Transplantation, Tongji Hospital, Tongji Medical College, Huazhong University of Science and Technology, Wuhan, Hubei 430030, China. Khodor I. Abou-Daya (  kha17@pitt.edu ) Thomas E. Starzl Transplantation Institute, University of Pittsburgh, School of Medicine, Pittsburgh, PA 15261, USA.


Introduction
Development and applications of the protocol Organ transplantation is a lifesaving treatment for patients with end-stage organ failure. Mouse kidney transplantation models have been widely used as an important investigational approach to study rejection 1,2 . Since the rst mouse kidney transplantation experimental model was established in 1973 3 , the techniques have been re ned and new techniques were developed to improve the survival rate of recipients 4,5,6,7,8,9,10,11,12,13,14 . In our lab, we continued to improve and utilize this model to study innate and adaptive mechanisms of rejection 15,16,17,18,19,20 . In the past three decades, signi cant advances have been made in prevention of acute graft rejection and improving early graft survival.
However, chronic graft rejection remains a major obstacle to long-term allograft survival 21 . It is therefore necessary to further our understanding of the pathophysiology of chronic rejection.
Graft rejection is caused by host immune response to non-self 22,23 . This response includes a circulating systemic compartment and a resident non circulating one. The resident compartment is maintained locally within the graft and most likely contributes to chronic rejection. The recently identi ed subset of non-circulating, tissue-resident memory T cells (T RM ) represents one important form of local immune responders within the grafts 24 . Another important contributor to the pathogenesis of chronic rejection is the formation of tertiary lymphoid organs (TLO) within the graft. TLOs represent important local immune response sites 25,26 . The functions of T RM in graft rejection are not known. The mechanism of TLO formation in the graft is also not clear. The inability to distinguish resident from host circulating systemic responses after transplantation remains an obstacle to studying local immune responses. Retransplantation of a graft to a congenically disparate secondary recipient offers the ability to discern and speci cally study the resident compartment of the primary recipient's immune response.
Re-transplantation of mouse heart allograft has been developed in our lab 27 and by other investigators 28 .
However, without treatment, heterotopic heart allografts are acutely rejected and are not life supporting 1 thus minimizing the utility of retransplanting a heart in the study of chronic rejection. Till date, mouse kidney allograft re-transplantation has not been reported primarily due to di culties in the secondary revascularization of the graft, re-establishment of the urinary tract continuity, and dissecting extensive adhesions during harvesting a transplanted kidney and ureter. We recently successfully established a mouse kidney re-transplantation model to investigate the role of T RM in kidney allograft rejection 19 . This detailed stepwise protocol will be of great value to nephrologists, immunologists, transplantation scientists, and microsurgeons interested in studying the local immune response.
Overview of the protocol: advantages and limitations Re-transplantation of a mouse kidney allograft includes ve stages: 1) primary donor graft harvest, 2) back-table preparation of the primary donor graft, 3) primary kidney transplantation, 4) secondary donor graft harvest from a primary recipient, and 5) re-transplantation of the secondary donor graft to a secondary recipient. We provide modi cations to previously reported mouse kidney transplantation procedures 5,6,8 in Stage 1 and 3 to improve survival. Despite its importance, Stage 2 has never been as well described. Stages 4 and 5 are novel and described in detail in this protocol.
Similar to most reported protocols, we opted to intra-abdominally position the mouse kidney graft at the recipient's right ank instead of ectopically implanting it in the neck 29 . Positioning the kidney at the ank is the most physiological and clinically relevant way. The kidney can be positioned at either ank. However, since the recipient's inferior vena cava (IVC) is closer to the right side, using the right ank will cause less tension at the venous anastomosis site and hence offer a decreased risk of venous stricture and bleeding. Left donor kidneys were preferred owing to the greater length of the left donor renal vein 3,4,5,6,7,8,9,10,11,12,13,14 . However, the right donor kidneys can also be transplanted 30 . To re-establish arterial blood supply in the primary transplantation procedure, we used end-to-side vascular anastomosis by suturing a segment of donor AO to recipient's AO. In contrast, the renal vein, a larger caliber vessel compared to the renal artery, was end to side anastomosed to the recipient's IVC. In the re-transplantation procedures, due to the di culty in dissecting the renal vessels of a previously transplanted graft, we opted to utilize segments of the secondary donor/primary recipient's AO and IVC for end-to side vascular anastomosis to secondary recipient's AO and IVC, respectively. In this protocol, we introduce an interrupted "6-stitch" suturing style for arterial anastomosis. We gained 100% vascular anastomosis success rate (last consecutive 100 operations).
For ureteric reconstruction, due to the risk of bladder patch necrosis and subsequent urine leak, we preferred to use ureteric implantation with our modi ed drag-in technique. However, since ureteric implantation and bladder patch techniques have both advantages and disadvantages 31, 32 , surgeons might opt to use either method depending on their experience. Details on how to effectively apply both methods are provided. Recipients underwent bilateral nephrectomy thus rendering the kidney grafts lifesupporting after transplantation. This allows surgical success to be assessed immediately after surgery by graft function and recipient's survival. Kidney function can be monitored by serum creatinine or blood urea nitrogen measurement.
To note, re-transplantation of heart or kidney allografts in rats had been used previously to study the in uences of early episodes after transplantation on the long-term graft outcome 33, 34, 35, 36 . However, although technically demanding, mouse kidney re-transplantation provides unique advantages over rat models, or other large animal models, because the mouse gene background is well characterized, the diversity and availability of transgenic mice are greater, and the animals and related research reagents are at a relatively lower price. Furthermore, the rejection of a mouse kidney allograft is slow 37 . This makes mouse kidney allograft re-transplantation a better model to study local immune responses, which need su cient time to be established after transplantation 19 . The purpose of this protocol is to provide repeatable stepwise procedures for mouse kidney re-transplantation.

Alternative Methods
Some microsurgeons adopt techniques that involve anastomosing cuffs of donor AO and IVC, similar to a carrel patch that is applied in clinical kidney transplantation, to the recipient AO and IVC, respectively 3,38 . Others use a heel-and-toe AO cuff for suture anastomosis 12, 39 and a cuff for non-suture renal vein connection 10 . These techniques are time consuming because the vascular patches/cuffs need to be prepared. Using segments of the donor AO and IVC instead of patches is a quicker 4 and a more widely used method. The donor AO segments can be superior renal 4,6,9,30,40 or inferior renal 5,8,30, 31 based on individual preference and surgical needs. Running sutures is widely adopted for both arterial and venous anastomosis. However, interrupted suturing of the renal artery has a higher success rate 31 .
For urinary tract reconstruction, four approaches are available: 1) Anastomosing donor bladder patch to a cystectomy on the recipient bladder dome in one-layer 3,4,5,8,38 , 2) Suturing donor bladder patch to the recipient bladder dome in two-layers with 7 or without 13 anti-re ux bladder seromuscular tunnel (similar to the Lich-Greoir technique), 3) Dragging donor ureter distal end into the recipient bladder 6,9,11,30,31,40 , and 4) Pulling the distal part of the ureter including a small bladder patch (2mmAE) into the recipient bladder 14 . Given the small size of a mouse's ureter, using a donor bladder patch was previously considered as the only practical choice for re-establishment of the ureteric continuity in mouse kidney transplantation 5 . After Han et al. introduced the ureter-dragging-into-bladder technique of ureteric implantation in 1999 6 , a growing number of investigators started to recognize its simplicity, feasibility, and superiority 9,11,30,31,40 . The latter technique also facilitates kidney re-transplantation 30   All microsurgical equipments must be sterile prior to utilization. Our instruments were autoclaved (>120°C for 30 min).

MICE
Authors recommend that all mice weigh 25-30 grams and are 8 to 12 weeks of age (The Jackson Laboratory).! CAUTION All experiments involving animals must be performed in accordance with national and institutional regulations. This protocol has been approved by the University of Pittsburgh and in accordance with criteria outlined in the Guide for the Care and Use of Laboratory Animals, a publication of the US National Institutes of Health. solely studying the local immune response, we considered mouse strains that lack secondary lymphoid organs including aly/aly and splenectomized B6.ltbr−/− mice or mice that lack an adaptive immune system such as B6.Rag−/−γc−/− mice. In the latter, lymphopenia induced proliferation might pose a caveat for studying local immunity. We opted for a splenectomized B6.ltbr−/−mouse.

Abdominal Wall Retractor (AWR) setup
Straighten the second loop of two 3 cm paper clips (Fig. 1e). Next, using mosquito forceps, bend the rst loop (wider) to make an 80-degree angle thus forming an abdominal hook. The third loop (narrower) is anchored to the cork part of the operating table using a 27-gauge needle. Each arm of the AWR is used to retract either left or right side of the abdominal wall.

Procedure
Primary Donor Surgery •TIMING 20-30 min 1│ Anesthesia. Place the primary donor in an induction chamber connected to an iso urane machine. Adjust the iso urane vaporizer to a maximum of 5% with continuous oxygen ow of 1 L/min. Anesthesia induction is veri ed by observing a decrease in mouse respiratory rate down to 1 breath per second.
2│ Skin preparation. Move the animal out of the induction chamber. Shave the abdominal skin entirely using an electric hair clipper. Alternatively, razor blades, depilatory creams or scissors can be used.
! CAUTION Make sure that the mouse is fully anesthetized to avoid accidental injuries to the limbs due to mouse's movements. Repeat step 1 if mouse is awake or awakening during skin preparation.
3│ Surgical Setup. Drape the surgical table in a sterile fashion. Place the anesthetized mouse on the thermostatic operating table under the surgical microscope. Set the vaporizer at 1-1.5% iso urane with continuous oxygen ow of 1 L/min for maintaining the mouse under general anesthesia using continuous iso urane inhalation through a mouse anesthesia mask. Tape all limbs to maintain the animal in the proper operative position on the operating table (Fig. 1a). Swab the abdominal wall with povidone-iodine rst and then with 70% ethanol.
! CAUTION Maintaining animal body temperature by using a thermostatic operating table or a heating pad during surgery is very important, especially for the recipients. Hypothermia will decrease the respiratory rate of the animal during surgery, signi cantly prolong the recovery time, and increase the mortality risk of the recipients after surgery. We highly recommend using a thermostatic operating table with a su cient cork area. Such a table is better for maintaining temperature, easier positioning, and anchoring the AWR during surgery.
4│ Make a midline abdominal incision from the xiphoid process to the pubis. After making the incision, decrease iso urane to 0.5-1% for maintenance of anesthesia.
▲CRITICAL STEP It is critical to observe the respiration of the mouse during the operation to assess the depth of anesthesia. Adjust the iso urane vaporizer to maintain a respiratory rate at approximately one breath per second. Deep anesthesia may lead to postoperative coma and death.
5│Using the AWR to retract the abdominal walls, expose the abdominal cavity. Anchor the retractors as speci ed in AWR setup section above (Fig. 1a). If the operating table lacks a cork component, corks or  styrofoam squares can be attached onto the operating table. 6│Make a sterile surgical wound drape by cutting a diamond hole (2×2 cm) in the middle of a dry nonwoven sponge (4×4 in). Cover the mouse with the drape as to only expose the abdominal surgical eld (Fig. 1a-c).
7│Wrap the small and large intestines using a warm wet nonwoven sponge (4×2 in) with the assistance of a wet cotton swab and place them on the cranial right side of the abdominal cavity to e ciently expose the left kidney for the subsequent procurement (Fig. 1a). ▲CRITICAL STEP Special attention should be given to protect the ureter's blood supply. Any damage to the ureter blood supply will increase the risk of ureter necrosis after surgery. The arrow in Fig. 3b shows the ureter accompanying vessels are carefully dissected and protected. To note, we use gonadal as a collective term for the internal spermatic and ovarian vessels in males and females, respectively.
10│ Dissect the abdominal aorta (AO) using the dumostar tweezers and micro forceps and place one segment of a 6-0 silk tie under the dissected portion of the AO between the levels of origin of left and right renal arteries (Fig. 2c).
! CAUTION Do not damage the lumbar veins draining into the IVC. Alternative AO levels where the silk tie is placed are used when the right renal artery or the superior mesenteric artery (SMA) origins are very close to the left renal artery origin. Under such circumstances, the silk tie should be placed above the level of the SMA origin and the ligations of right renal artery and SMA will be performed during the back-table procedures using 10-0 nylon.
11│ Gently dissect the portion where the left RV drains into the IVC using the dumostar tweezers and micro forceps to separate it from the anterior surface of the AO. Create a space under the left RV that allows the tip of the closed arms of the micro forceps to completely pass beneath it (Fig. 3c).
▲CRITICAL STEP The pre-prepared space guarantees that the RV can be immediately transected once the AO is ligated (step 13). This provides earlier graft perfusion and reduces graft ischemia reperfusion (IR) time.
▲CRITICAL STEP To avoid a potential air embolus, remove air bubbles prior to the infusion.
13│ Ligate the AO using the silk suture that has already been placed under the AO at step 10 ( Fig. 2c). By stretching the scissor blade through the space made in step 11, immediately cut the left RV at the level where it drains into the IVC using the corneal scissors. This will serve as a perfusion outlet (Fig. 2d). Then, immediately start step 14. After cutting the RV, the mouse will bleed out. Cutting the AO in step 17 will further bleed the mouse and accelerate cardiac arrest. At such time, the anesthesia machine can be turned off.
▲CRITICAL STEP The right gonadal artery (Fig. 2i) will be cut in most mice while cutting the left RV and will be ligated during the back-table procedures. We do not recommend doing any dissection or ligation to the right gonadal artery branches before cutting the left RV, because it is easier and quicker to perform this during the back-table procedures after the kidney graft is procured and will avoid any injuries to the vessels due to extra dissections at this step.
14│ Clamp the AO approximately 1 cm under the origin of the left renal artery. Slightly lift the AO up using the dumostar tweezer. Slowly inject 4 ℃ of cold heparinized saline solution prepared in step 12 by puncturing the AO just above the clamp site (Fig. 2e). When blood from the donor kidney is ushed out and the kidney turns evenly pale, stop the perfusion. It usually requires 0.5-1 ml of perfusion solution to achieve adequate perfusion of the graft.
15│ Cut the AO 5 mm below the left renal artery origin from the AO using the corneal scissors (Fig. 2f).
16│ Cut the IVC, connective tissue, and all lumbar arterial branches along the right side of the AO till the level of the silk ligation which has been already performed in step 13.
▲CRITICAL STEP Make sure not to cut the lumbar arterial branches (Fig. 2i) too close to the AO. Leave enough length for an easier ligation during the back-table procedures.
17│ Cut the AO above the ligation which has been already performed in step 13 (Fig. 2g).
18│Use a dry nonwoven sponge (4×2 in) to absorb the blood and perform a right cranial visceral rotation. Better visualization of the AO segment and renal pedicle will be achieved.
19│ Separate the connective tissues between the aortic segment and renal artery from the posterior wall of the abdominal cavity.
20│ Transect the left ureter at the distal end (Fig. 2h). Separate it from the surface of the psoas major muscle.
▲CRITICAL STEP The ureter transection should be placed as distal as possible so that the ureter length is su cient for re-transplantation. ▲CRITICAL STEP Careful examination of the AO and RV ends is needed to make sure they were evenly cut and hence will form adequate anastomoses.
25│Perfuse the organ through the AO using 0.2-0.5 ml 4 ℃ cold heparin saline in a 27-gauge syringe to eliminate any residual blood inside the graft and vascular lumens. Ligate the right gonadal artery which originates from the anterior wall of the AO and the lumbar arteries which originate from the right or posterior wall of the AO using 10-0 nylon (Fig. 2i). Ligate the right renal artery or SMA using 10-0 nylon if alternative AO ligating levels as described in step 10 are applied.
▲CRITICAL STEP In our experience, most mice's right gonadal arteries originate from the AO anterior wall and are more cranial than the draining site of the left renal vein into the IVC. Also, almost 50% of the mice have 1-2 lumbar arteries originating from the right or posterior wall of the AO portion which need to be harvested as a segment attaching to the renal artery. These arterial branches are small and hard to expose for ligation before perfusion (Fig. 2i). We highly recommend that the ligations of these branches are performed during back- 28│Wrap the small and large intestines using a warm wet nonwoven sponge (4×2 in) with the assistance of a wet cotton swab and perform a left cranial visceral rotation to e ciently expose the right abdominal cavity and recipient IVC and AO (Fig. 1b).
29│ Slightly lift the right ureter using the dumostar tweezers and cut the ureter with a mono-polar ne-tip cautery. Ligate the native right renal artery and vein together using 6-0 silk tie and remove the right native kidney of primary recipient (Fig. 4a).
30│ Use 6-0 silk to temporarily ligate the small lumbar arteries and veins that originate from and merge into the AO and IVC respectively to prepare a portion of the AO and IVC for anastomosis. Dissect and remove the fat and connective tissue that adhere to the anterior walls of the AO and IVC.
▲CRITICAL STEP There is no need to separate the AO from the IVC. The ideal position for arterial and venous anastomosis is between the origin of left RV and the iliac bifurcation of AO (the anastomosis window). The left or right lumbar veins usually drain into the renal veins. If they drain into the IVC, temporarily ligating them at the IVC may offer better preparation of AO and IVC as ideal anastomosis sites.
▲CRITICAL STEP Multiple temporary ligations might be needed to achieve an ideal AO anastomosis window, although usually 1-2 temporary ligations are su cient. More than 2 temporary ligations are often used when there is a horizontal arterial branch originating from the AO running across the anterior wall of the IVC. In this case, longer portions of AO and IVC need to be prepared for the anastomosis. In addition, the anastomosis site might need to be moved to the upper or lower part of the anastomosis window so that the arterial branches can be avoided. However, if there is no way to avoid them, the branches need to be ligated.
31│Place the rst bull-dog clamp on the distal part of the IVC and AO using the micro forceps and mosquito forceps holding the bulldog (Fig. 5).
32│Place the second bull-dog clamp on the proximal part of the IVC and AO similar to step 31 (Fig. 5).
▲CRITICAL STEP The distance between the two bull-dog clamps should not be less than 5 mm. It is necessary to gently press the IVC anterior wall toward the cranial direction using the micro forceps before nally placing the second bull-dog clamp so that the IVC is not overly dilated. Over dilation of the IVC will affect the operative exposure of the AO. It is also critical to observe the distension of the IVC between the clamps after the second bull-dog clamp is placed. If the distention of the clamped portion of the IVC continues, this represents unclamped venous in ow to the IVC and the temporary ligations placed in step 30 should be examined and adjusted to prevent any blood ow into the clamped part of the IVC. If not prevented, this will severely affect the subsequent anastomosis procedure and can be fatal to the recipient. See Supplementary Video 2 for additional guidance on placing temporary ligations and bulldog clamps to prepare recipient AO and IVC for anastomosis. 33│Make a stitch at the anterior wall of the AO using the micro forceps holding 10-0 needle and leave the needle on the AO wall. Create an arteriotomy at the anterior wall of the AO by cutting the anterior wall under the bottom of the needle that stays on the AO wall using the vannas spring scissors (Fig. 5a).?

TROUBLE SHOOTING
▲CRITICAL STEP The recipient aortic arteriotomy should match the lumen size of the graft donor AO.
Cutting too much of the anterior wall of the recipient AO will lead to a size mismatch between donor AO lumen and recipient AO arteriotomy. This will also increase the risk of stricture at the arterial anastomosis site and is fatal to the recipient. 34│Flush the AO through the opening at the anterior wall immediately using warm non-heparinized normal saline in 1 ml syringe with 27-gauge needle. Stop ushing when all blood and any clogs are ushed out of the portion of the AO between clamps and the portion turns completely pale. Usually, it requires 0.3-0.5 ml saline to achieve an adequate ush of the AO.
▲CRITICAL STEP Do not touch the AO intima with the syringe needle during ushing. Any damage to AO intima will increase the risk of thrombogenesis at the arterial anastomosis site. While ushing, directed cranially, gently and repetitively squeeze the AO anterior wall caudal to the anastomosis site with micro forceps. It is unnecessary to ush caudally.
▲CRITICAL STEP Use the non-heparinized normal saline instead of heparinized normal saline to ush the AO before and during the anastomosis procedures to avoid prolonged bleeding at the anastomosis site after releasing the bulldogs.
35│Gently place the primary donor kidney on the right ank side of the recipient abdominal cavity. Use the micro forceps to hold the 10-0 needle with nylon and end-to-side anastomose the donor AO at the prepared site of the recipient AO. The anastomosis is established using the "6-stitch" technique using interrupted sutures with 10-0 nylon (Fig. 4b). Brie y, make the rst stay stitch at 12 o'clock and the second stay stitch at 6 o'clock to x the donor AO to the opening of the recipient AO (Fig. 5b). Secondly, make the third and fourth stitches at 2 o'clock and 4 o'clock (Fig. 5c). Then, rotate the operating table by 180 degrees and gently ip the kidney graft over to the left side of the recipient abdominal cavity (Fig.   1c). Flush the anastomosis site with normal saline. Make the fth and sixth stitches at 2 o'clock and 4 o'clock (8 o'clock and 10 o'clock, respectively, before table rotation) to nish the arterial anastomosis (Fig.   5d).
▲CRITICAL STEP Make sure that full thickness passes of the suture needle including the vascular adventitia and the intima are achieved and no posterior AO wall is hooked when suturing the anterior wall of the AO.
▲CRITICAL STEP Examine the 10-0 needle prior to suturing the AO wall. Any damage of the tip of the needle will increase the perforating injury to the AO wall and the potential to carry the adventitia into the lumen of the AO. This increases the risk of arterial thrombogenesis at the anastomosis site.
▲CRITICAL STEP Employ interrupted suturing ("6-stitch" method) instead of a continuous running suture for arterial anastomoses, which guarantees arterial potency without stenosis or thrombosis after unclamping the AO. We have found that using more than 6 stitches is unnecessary. See Supplementary Video 4 in re-transplantation for additional guidance on the "6-stitch" technique of arterial anastomosis, which works in both primary transplantation and re-transplantation procedures.
36│After nishing the arterial anastomosis, keep the recipient and the kidney graft in the same position for venous anastomosis. Use the Vannas-Tübingen spring scissors to make a venotomy at the anterior wall of the IVC with the assistance of micro forceps (Fig. 5e). The primary donor renal vein is end-to-side anastomosed to the recipient IVC using a continuous running suture with 10-0 nylon (Fig. 4c).? TROUBLE SHOOTING First, place two stay stitches at 12 o'clock and 6 o'clock, respectively, to x the primary donor RV end to the anastomosis site of the IVC. Second, nish the anastomosis of posterior wall inside the IVC lumen from 6 o'clock to 12 o'clock and let the needle go out of the lumen at 12 o'clock ( Fig. 5f-g). Last, nish the anastomosis of the anterior wall from 12 o'clock to 6 o'clock. Flush the IVC with warm normal saline to expel any air bubbles in the IVC before nishing the last stitch (Fig. h).
▲CRITICAL STEP Flush all air bubbles out of the IVC before nishing the venous anastomosis. Any air bubbles in the IVC will lead to air embolus and may be fatal.
37│Gently ip the kidney back to the right ank side of the recipient. Slightly press the arterial anastomosis site with a dry cotton swab and remove the caudal bulldog clamp rst and then release the cranial clamp using the mosquito forceps. Keep the swab on the arterial anastomosis site with gentle pressure for 10 seconds to stop the bleeding then slowly remove.
▲CRITICAL STEP Gelatin sponge can be applied to the anastomosis site for hemostasis.
38│Rotate the operating table again by 180 degrees so that the animal caudal end faces the surgeon. 39│Remove all temporary ligations placed in step 30.
40│Double check the anastomosis sites of the AO and IVC, and the graft pedicle. Obtain hemostasis by using dry cotton swab, gelatin sponge, cautery, or ligation. Few recipients need the above measures and most bleeds can be resolved by applying gentle pressure with swab, or cautery. See Supplementary Video 5 in re-transplantation for additional guidance on venous anastomosis, releasing the bulldogs, achieving hemostasis at the anastomosis site, and removing the temporary ligations, which works in both primary transplantation (RV to IVC, AO to AO) and re-transplantation (IVC to IVC, AO to AO) procedures.
41│Separate the connective tissues and fats from the distal part of the ureter graft using the dumostar tweezers and Vannas-Tübingen spring scissors (Fig. 6a).
▲CRITICAL STEP Do not dissect the ureter too much to avoid injuring the ureter's arterial supply. Examine the ureter under the microscope and make sure the blood vessels are intact. The distal ureteral part that lacks its arterial supply will be transected off later.
42│Ligate the end of the ureter graft with 10-0 nylon to let the ureter dilate (Fig. 6a).
▲CRITICAL STEP The dilation of the ureter at this step will help avoid ureter torsion and twisting during the ureter to bladder implantation. 43│Puncture through the recipient bladder from the anterior wall to the posterior wall using the Dumostar tweezers with assistance of Dumont forceps holding the bladder top.
▲CRITICAL STEP Do not use the needle to puncture the bladder. Puncturing with a blunt device (tweezers' tip) offers better healing of the bladder seromuscular layer and decreases the risk of a urine leak. 44│Hold the 10-0 nylon ligated at the end of the ureter with the Dumostar tweezers and slowly pull the ureter through the recipient bladder from the posterior wall to the anterior wall (Fig. 6b).
▲CRITICAL STEP Do not grab the ureter directly. Guiding the ureter through the bladder by grabbing the 10-0 nylon pre-ligated at the ureter end will decrease injuries to the ureter. It also provides a chance to reguide the ureter through or out of the bladder before appropriately xing the ureter onto the posterior wall of the bladder. 45│Make two stitches using 10-0 nylon to x the connective tissues that accompany the ureter to the posterior seromuscular wall of the recipient bladder (Fig. 4d, 6d).
▲CRITICAL STEP Two symmetrically distributed stitches should be made exactly at the same level of the ureter. They should be placed at the edge of the opening in the posterior wall formed by puncturing the bladder with the tweezers (Fig. 6d).
46│Pull the ureter out of the anterior side of the recipient bladder using the 10-0 nylon ligature and cut the distal part of the ureter. Gently lift the anterior wall of the bladder and let the end of ureter graft retract spontaneously into the recipient bladder (Fig. 6c).? TROUBLE SHOOTING 47│Make a " gure of eight" stitch to close the hole at the anterior wall of the recipient bladder (Fig. 4e,   6c).
An alternative approach to establish the primary donor ureter to primary recipient bladder connection is attaching donor ureter with a donor bladder patch to the recipient bladder dome. When using this technique, the whole donor bladder with posterior urethra and whole left donor ureter should be procured en bloc (Fig. 7).
▲CRITICAL STEP For the alternative technique, make sure the donor bladder patch is adequately nourished (Fig.7a, c and d) otherwise (Fig.7b) techniques described in step 41-46 should be adopted. The donor bladder patch should be trimmed down to an adequate size. A small sized patch (Fig.7c-d) risks ureteric obstruction caused by inappropriately suturing the ureteral ori ce on the donor patch. Larger patches need more blood supply and inadequate blood supply will cause patch necrosis after surgery. The opening at the recipient bladder dome should also be as small as possible. It can be dilated with micro forceps to the size that matches the donor bladder patch. If the donor bladder patch is too large (and not necrotic), it will likely cause urine retention due to neurogenic bladder, which is fatal. The 9-0 absorbable vicryl sutures are recommended for suturing the donor bladder patch to the recipient bladder dome. In our experience, the technique of suturing donor bladder patch to recipient bladder dome has no apparent advantages when compared with dragging the donor ureter into the recipient bladder as described above, especially when using Balb/c mice as donors. Balb/c donor bladder patch has high risk of necrosis after surgery likely due to insu cient blood supply (Fig.7b). We recommend using the ureteric implantation (ureter-drag-in) technique to establish ureter-bladder connection in this model. See Supplementary Video 6 in re-transplantation for additional guidance on the ureteric implantation procedure.
48│ Displace the small and large intestines to the right side of the recipient using a wet cotton swab. Cauterize the recipient left ureter (Fig. 4f). Ligate the recipient left renal artery and vein together using 6-0 silk (Fig. 4g) and remove the native left kidney.
49│ Restore the small and large intestines to the recipient abdominal cavity and carefully re-position them using a wet cotton swab. 50│ Leave 0.5 ml warm normal saline in the abdominal cavity and remove the wound drape and the retractors. 53│Observe the recipient mouse every 5 min until it is fully awake. Recipient mouse recovers within 20 min. Mouse is given ad libitum access to water and an antibiotic (Trimethoprim 275 ppm, Sulfadiazine 1365 ppm) chow for three weeks after surgery. After three weeks, chow is switched to regular. Secondary Donor Surgery•TIMING 40-60 min 54│ Anesthesia and laparotomy. Anesthetize the primary recipient mouse, clean the abdominal hair, prep the skin, make a midline incision, retract the abdominal wall to expose the abdominal cavity, and cover the mouse with pre-cut wound drape similar to steps 1-6 in primary donor surgery.
55│ Wrap the small and large intestines and place them on the left side of the mouse to expose the kidney graft (Fig. 1b).
56│ Use a ne-tip cautery to dissect the kidney and ureter grafts, the kidney graft's artery and vein, and the mouse's bladder, AO and IVC. Separate them from the adherent connective tissues, intestines, and liver tissues. Separate the kidney and ureter from the posterior wall of the abdominal cavity.? TROUBLE SHOOTING 57│ Ligate and transect all lumbar arteries and veins that respectively originate from and drain into the AO and IVC from 1 cm below and 0.5 cm above the graft anastomoses. Skeletonize this portion of AO and IVC together and separate them from the posterior abdominal wall (Fig. 8a). There is no need to separate the IVC from the AO.? TROUBLE SHOOTING 58│ Ligate the AO and IVC together above the level of the graft anastomosis site using 6-0 silk. Make a venotomy on the IVC at the bifurcation level immediately to make an outlet for the perfusion solution. Slightly lift the AO by clamping it above the bifurcation level using dumostar tweezers. Slowly inject 50 U/ml cold 4 ℃ heparin saline in 1 ml syringe with a 27-gauge needle to perfuse the secondary donor kidney graft. The needle should be inserted slightly below >1 cm where the primary donor kidney is anastomosed but above the AO clamp site (Fig. 8b-d). When the blood from the kidney graft is ushed out and the kidney graft turns evenly pale, stop the infusion. It usually requires 0.5-1 ml of perfusion solution to achieve adequate perfusion of the graft. 59│ Transect the AO and IVC together at the level of 0.5 cm below the anastomosis site of the kidney graft using corneal scissors (Fig.8e).
60│ Transect the AO and IVC together above the silk ligation performed in step 58 (Fig. 8f).
61│ Transect the ureter graft at its entry into the bladder (Fig. 8g). Immediately harvest the kidney graft with connections of whole ureter, renal vein, renal artery, primary donor AO, and skeletonized secondary donor AO and IVC. Put them in 4 ℃ cold normal saline for preservation.
62│Euthanize the secondary donor according to institutional guidelines. See Supplementary Video 3 for additional guidance on procuring transplanted kidney graft for retransplantation.
Secondary recipient surgery •TIMING 50-60 min 63│ Re-transplant the kidney graft from the secondary donor to the secondary recipient by repeating steps 27-51. In comparison to the primary recipient surgery, the differences in secondary recipient surgery for kidney graft re-transplantation include: 1) Instead of using the primary donor mouse's AO for anastomosis, the graft renal artery with a segment of the primary donor mouse's AO and a segment of the primary recipient mouse's AO is end-to-side anastomosed to the secondary recipient mouse's AO (Fig. 9b) See Supplementary video 4 for additional guidance on arterial anastomosis during re-transplantation; 2) Instead of using renal vein for anastomosis, the graft renal vein with a segment of the primary recipient's IVC is end-to-side anastomosed to the IVC of the secondary recipient mouse (Fig. 9c). See Supplementary video 5 for additional guidance on venous anastomosis during re-transplantation; 3) In step 45, instead of 2 stitches, a total of 3-4 well-proportioned stitches are needed to x the ureter graft onto the posterior wall of the secondary recipient mouse's bladder due to thickening of ureter graft's wall after primary transplantation (Fig. 9d, d-2). See Supplementary video 6 for additional guidance on establishing connection and xation of the ureter graft to the bladder of the secondary recipient during retransplantation.? TROUBLE SHOOTING Secondary recipient recovery•TIMING 15-20 min 64│Secondary recipient mouse was treated similarly to the primary recipient mouse. 65│Secondary recipient mouse recovers similarly to the primary recipient mouse within 20 min after surgery.

Anticipated Results
The mouse kidney re-transplantation model has not been previously reported. We have achieved >90% success (close to 100% for vascular anastomoses) in mouse primary kidney transplantation prior to employing mouse kidney re-transplantation. In total, 6 allogeneic (F1.OVA kidney to B6.CD45.1 to B6.ltbr-/-) and 2 syngeneic (B6 to B6 to B6) re-transplantations were completed. The syngeneic and allogeneic re-transplants were performed on 14 d and 35 d, respectively, after primary kidney transplantation. The pilot allogeneic re-transplant recipient survived 11 d post re-transplant. 4 out of 5 subsequent allogeneic recipients survived until the set endpoint and were harvested 70 days post retransplantation. One recipient died at 21 days. 2 syngeneic recipients survived till set harvest date of 30 days after re-transplantation. All recipients' grafts were functional post re-transplant with normal serum creatinine of < 0.2 mg/dl at harvest or prior to unanticipated death. Hematoxylin and eosin-stained sections of syngeneic primary or re-transplanted kidney grafts obtained 30 d after primary transplantation or re-transplantation is shown in Figure 10. The syngeneic primary and secondary grafts' histological sections showed no pathology.       Experiments were performed under an institutional animal care and use committee-approved protocol.