Orthotopic Kidney Auto-Transplantation in a Porcine Model Using 24 Hours Organ Preservation And Continuous Telemetry

Summary

Large animal models play an essential role in preclinical transplantation research. Due to its similarities to the clinical setup, the porcine model of orthotopic kidney auto-transplantation described in this article provides an excellent in vivo setting for the testing of organ preservation techniques and therapeutic interventions.

Abstract

In the present era of organ transplantation with critical organ shortage, various strategies are employed to expand the pool of available allografts for kidney transplantation (KT). Even though, the use of allografts from extended criteria donors (ECD) could partially ease the shortage of organ donors, ECD organs carry a potentially higher risk for inferior outcomes and postoperative complications. Dynamic organ preservation techniques, modulation of ischemia-reperfusion and preservation injury, and allograft therapies are in the spotlight of scientific interest in an effort to improve allograft utilization and patient outcomes in KT.

Preclinical animal experiments are playing an essential role in translational research, especially in the medical device and drug development. The major advantage of the porcine orthotopic auto-transplantation model over ex vivo or small animal studies lies within the surgical-anatomical and physiological similarities to the clinical setting. This allows the investigation of new therapeutic methods and techniques and ensures a facilitated clinical translation of the findings. This protocol provides a comprehensive and problem-oriented description of the porcine orthotopic kidney auto-transplantation model, using a preservation time of 24 hours and telemetry monitoring. The combination of sophisticated surgical techniques with highly standardized and state-of-the-art methods of anesthesia, animal housing, perioperative follow up, and monitoring ensure the reproducibility and success of this model.

Introduction

Since the first successful human renal transplantation between identical twins in 1954, performed by the pioneering group of the Nobel prize laureate surgeon Joseph Murray¹, kidney transplantation (KT) has evolved as the mainstay of treatment for patients with end-stage renal disease (ESRD)². KT shows superior long-term clinical outcomes and quality of life compared to dialysis². Short- and long-term survival rates after KT improved continuously, due to advances in surgical techniques, organ preservation, immunosuppressive therapy, and critical care, hence KT became widely available on a global scale^2,3,4.

Due to critical organ shortage, there is a continuously increasing gap between allograft supply and demand^3,5,6. In 2018, approximately 12,031 patients were waiting for KT in Germany, however, only less than 20% (2,291 patients) could receive a donor kidney due to the extreme shortage in organs for transplantation⁷. Unfortunately, not only the absolute number of organ donors, but also the general quality of the allografts offered for transplantation have declined in the past decades^8,9. An increasing tendency was observed in the numbers of predamaged or "marginal" kidney allografts that had to be accepted for transplantation¹⁰. The use of ECD allografts may reduce waiting time and waiting list morbidity and mortality, it is, however, associated with an increased incidence of graft-related complications such as primary graft non-function (PNF) and/or delayed graft function (DGF)^8,9,10. Further research is essential to optimize allograft utilization, expand the donor pool and protect and recondition marginal allografts which ultimately may improve patient outcomes^3,6.

Due to the resource-intensive and complex nature of large animal transplantation models, a large number of studies are performed using small animals or in ex vivo settings^{11,12,13,14,15}. Although these models can deliver important scientific data, the translation of these findings to the clinical setting is often limited. The porcine model of orthotopic kidney auto-transplantation is a well-established and reproducible model that allows to test new innovative treatment approaches in a clinically relevant in vivo setting, with potentially longer follow-up periods and abundant possibilities for repetitive sample collection^16,17. Beyond the advantage of the comparable size, which allows relatively direct translation into the clinical setting (particularly for medical device development and drug dosage), the surgical-anatomical and physiological similarities in terms of ischemia-reperfusion injury (IRI) response and kidney damage, support the use of this model in translational research^17,18,19. This model also provides an excellent training opportunity to prepare young transplant surgeons for the technical challenges of clinical organ transplantation²⁰.

There are also multiple differences compared to the human setting and various technical modifications of the model can be found in the literature^{16,17,19,20,21}. This article comprehensively describes technical details, pitfalls, and recommendations which can aid to establish the model of porcine orthotopic kidney auto-transplantation. The described telemetry and video monitoring method as well as our specifically designed housing facility allows a close-up severity assessment and clinical observation of the animals. The use of a percutaneous urinary catheter and designated porcine jackets provide the possibility of a detailed assessment of kidney function without the use of metabolic cages. These technical modifications are described as potential solutions to comply with the modern challenges of the 3R principle (Replacement, Reduction and Refinement) and improve animal experiments using large animal models²².

Protocol

The present study was designed according to the principles of the ARRIVE (Animal Research: Reporting of In Vivo Experiments) guidelines²³. Experiments were performed in accordance with the institutional guidelines and the German federal law regarding the protection of animals. The full ethical proposal was approved by the responsible authorities (Governmental Animal Care and Use Committee, LANUV NRW - "Landesamt für Natur, Umwelt und Verbraucherschutz Nordrhein-Westfalen", Recklinghausen, Germany, Protocol ID: 81-02.04.2018.A051). All animals in the present study received humane care according to the principles of the "Guide for the Care and Use of Laboratory Animals" (8th edition, NIH Publication, 2011, USA) and the Directive 2010/63/EU on the protection of animals used for scientific purposes (Official Journal of the European Union, 2010). Female German landrace pigs were obtained from a hygienically optimized barrier breeding facility (Heinrichs GbR, Heinsberg, Nordrhein-Westfalen). Figure 1 depicts the summary of the described experimental protocol.

1. Animals and housing

1. Use female German landrace pigs (or comparable) for this protocol. Deliver the animals to the research facility 14 days before the first surgery (telemetry implantation) for acclimatization and house them in a temperature- and humidity-controlled barrier environment with a 12 h light and dark cycle ( Figure 2).
2. Mount two telemetry receivers on the ceiling of the room which allows the registered data to be transferred directly to a PC located in the observation room. Ensure that animals are observed visually during the regular visits by the veterinary officers and by the animal caretaker in charge (every 8 h and on-demand).
   NOTE: Furthermore, in this experiment a real-time camera footage with integrated thermal imaging connected to the local network was used. Details of the housing facility used in this study are depicted in Figure 2.

2. Basic techniques and common procedures

1. Fast the animals overnight before surgery.
2. Premedicate by an initial intramuscular injection of azaperone (4 mg/kg) and atropine (0.1 mg/kg), followed by an injection of ketamine (15 mg/kg) 10 min later.
3. After premedication, weigh the animal and transfer it directly from the housing facility to the central OR facility anesthesia preparation room.
4. Cannulate one of the large ear veins using an 18 G peripheral venous catheter. Monitor the animal by a standard ECG and pulse oxymetry.
5. Initiate the anesthesia with propofol (3 mg/kg).
6. Expose the vocal cord with a laryngoscope and insert a 7.5 mm endotracheal tube. The cuff is blocked with air according to manufacturer´s recommendations.
7. Insert an oro-gastric drainage tube to remove fluid and air from the stomach.
8. Insert a urinary catheter via the urethra.
9. Subsequently, trim the skin in the area of the surgical incision.
10. Apply eye ointment to prevent drying of the cornea during surgery.
11. After orotracheal intubation, maintain anesthesia with isoflurane (final expiratory 1.45-2.0 Vol.%) and fentanyl (3 -7.5 µg/kg/h).
12. Ensure active intraoperative temperature control of the animal by a heating pad and using warmed air. Insert a rectal probe to monitor body temperature (target temperature 36.5 °C - 37.5 °C).
13. Administer antibiotic prophylaxis using cefuroxime (35 mg/kg i.v.). Infuse Ringer solution at 4 mL/kg/h and increase to 8 mL/kg/h after skin incision. Administer a prophylactic dose of pantoprazole (40 mg i.v.) over the ear vein access.
14. Perform all surgical procedures under sterile conditions according to the general principles of surgical asepsis and antisepsis. Disinfect the surgical field with povidone-iodine solution and cover with surgical drapes.

3. Telemetry implantation

1. Prepare the animal for surgery following the steps described under section 2 of the protocol and confirm proper anesthesia by a decreasing heart rate and a lack of conscious movement of the animal.
2. Collect blood and urine samples to determine individual baseline lab values.
3. Mark the incision sites using a permanent marker.
4. To implant the arterial sensor of the telemetry transponder, perform a 3-4 cm incision in the groin. Expose and dissect the artery in a 360˚ fashion.
5. Using an Overholt clamp pull through two-vessel loops below the artery and secure them with mosquito clamps.
6. After making an arteriotomy using #11 blade scalpel, insert the arterial sensor. Close the arteriotomy using 5-0 polypropylene suture with single knot sutures and secure the arterial sensor using one of these sutures.
7. Make a 3-4 cm large incision on the left flank of the animal and create a subcutaneous pouch for the transponder by blunt dissection.
8. Tunnel the telemetry transponder to the flank and fix it to the muscle fascia (3-0 polypropylene, single knot). Tunnel the red and white ECG electrodes to the right and left side of the thorax. Make two 1 cm incisions and secure the electrodes in the muscle tissue to ensure a good ECG signal with single knot sutures (3-0 polyglactin).
9. Commence registration of telemetry data and check the various signals (e.g., body temperature registered by the transponder body itself, arterial blood pressure, and ECG signals).
10. Close the incisions in the groin, on the left flank and the two small thoracal incisions using muscle and subcutaneous sutures (3-0 polyglactin) and close the skin using a non-absorbable monofilament suture (e.g., 2-0 Prolene).
11. Use a spray film dressing to seal the incision sites.
12. At this time point prepare the animal to wear a designated porcine jacket which the animal wears for the rest of the study period. Replace jackets with a clean jacket following every surgical intervention.
    NOTE: To record stable baseline data, telemetry transponders are implanted 14 days before the index surgery (left nephrectomy, please also see Discussion).

4. Nephrectomy and kidney graft retrieval

1. Prepare the animal for graft retrieval surgery following the procedures described in section 2.
2. After the induction of anesthesia, cannulate the external jugular vein. Following the sterile disinfection of the surgical field, a 4 cm incision is made on the right side in the jugular groove.
3. Dissect subcutaneous tissue and muscle to expose the external jugular vein.
4. Expose and dissect the vein in a 360˚ fashion.
5. Using an Overholt clamp pull through two-vessel loops below the vein and secure them with mosquito clamps.
6. Tunnel the jugular catheter to the back of the animal. For this, position the pig on its left side.Use the Seldinger method to insert the jugular catheter.
7. Close the opening on the vein and secure the catheter using 5-0 polypropylene suture.
8. Close the incision in two layers (e.g., 3-0 polyglactin for the muscle and subcutaneous and 2-0 polypropylene for the skin).
9. Secure the catheter to the skin with multiple sutures (2-0 polypropylene).
10. Test the jugular vein catheter for free aspiration and injection. Subsequently, switch the intravenous line from the ear vein cannula to the central venous line.
11. Following surgical disinfection and draping, perform a median laparotomy to open the abdomen (25-30 cm). Use a standard abdominal retractor to expose the surgical field.
12. Use wet and warm abdominal towels to cover the colon and the small bowel. Ask the second assistant to hold the bowel to the direction of right hemi-abdomen exposing the kidney and its vascular structures.
13. Open the peritoneal layer and dissect the left kidney and the ureter from any adherent tissue using the monopolar cautery, bipolar forceps, and fine scissors.
14. Ligate and divide the left ureter (3-0 polyglactin) leaving an at least 10 to 12 cm long segment.
15. Complete the dissection of the left renal vein(s) and artery to their origin from inferior vena cava and aorta, respectively.
    NOTE: Avoid injury and opening of the large lymphatic vessels in this anatomical region. Also be aware of a potential injury to the azygo-lumbar vein joining to the renal vein near to its origin from the vena cava.
16. Dissect and ligate the azygo-lumbar vein between two ligatures (3-0 polyglactin).
17. Prepare for the back-table dissection using a bowl of ice and a sterile cover.
18. To retrieve the graft kidney, clamp the renal artery and the renal vein close to the aorta and the vena cava with vascular clamps. Remove the kidney graft by cutting the vessels with a scissor close to the clamps then hand the kidney over to the back-table team.
19. Close the stump of the renal artery using a 5-0 polypropylene suture. Close the renal vein using a two-layer continuous suture with 5-0 polypropylene. Remove the vascular clamps.
20. After checking the area for bleeding or lymphatic leakage, close the abdomen in 4 layers.
    NOTE: Peritoneum: 3-0 polyglactin running suture; fascia: 0 polyglactin running suture; subcutaneous layer: 3-0 polyglactin running suture; skin: skin staplers after kidney retrieval surgery, to facilitate re-opening the abdomen the following day and 2-0 polypropylene single knot sutures after the transplantation procedure for definitive closure.
21. After applying sterile wound dressing, return the animal to the housing facility and allow to recover following endotracheal extubation. For postoperative analgesia, use buprenorphine (0.05 - 0.1 mg/kg) intramuscularly every 8 h until auto-transplantation.

5. Back-table and organ preservation

1. After graft retrieval, immediately cannulate the renal artery using a standard 14 G (orange) peripheral catheter and fix it using a tourniquet prepared from 3-0 polyglactin.
2. Rinse the kidney with cold organ preservation solution.
3. After flushing with 500 mL of organ preservation solution, remove the arterial cannula, wrap the kidney graft in sterile organ bags and store in organ preservation solution with a target cold ischemic time (CIT) of 24 h at 4 °C using a computer-controlled cooling circuit.
   NOTE: A brief post-preservation flush is recommended using 500 mL of 4 °C normal saline solution.

6. Contralateral nephrectomy and orthotopic kidney auto-transplantation

1. During the recipient operation, adapt premedication and initial anesthesia to the restricted renal metabolism and avoid the use of ketamine. Induction is performed with propofol (3-5 mg/kg i.v.), midazolam (0,25 mg/kg i.v.), and atropine (0.1 mg/kg i.m.). Thereafter, the preoperative preparation is identical to the procedures described in section 2.
2. Maintain anesthesia with isoflurane (final expiratory 1.45-2.0 Vol.%) and fentanyl (3 - 7,5 µg/kg/h) and propofol (2 - 4 mg/kg/h).
3. Check and continuously monitor ECG, pulse oximetry, rectal temperature and the function of the telemetry transponder.
   NOTE: Strict anesthesia and blood pressure control is of crucial importance during the implantation procedure.
4. In rare cases where the arterial blood pressure signal registered over the telemetry transponder is not satisfactory due to the supine position of the animal, place a further arterial catheter into the right femoral artery using percutaneous puncture and the Seldinger technique.
5. Following sterile draping, reopen the median laparotomy and expose the surgical field using the abdominal retractor. The colon and small bowel are placed to the left side of the abdomen to expose the intact right kidney.
6. Similar to the donor procedure, dissect the contra-lateral kidney and its vessels from the surrounding tissue. Dissect the right renal vein and renal artery in the direction of the kidney hilum to ensure sufficient vessel length for anastomosis.
7. 5 min before vascular clamping, inject natrium-heparin intravenously (100 I.U./ kg).
8. Clamp the right renal artery and the right renal vein using vascular clamps. The right kidney is removed. The vessels are checked for integrity before starting the anastomoses.
9. Place the preserved graft kidney into the abdomen and start the venous and arterial anastomoses.
10. From this point onwards, keep the mean arterial pressure over 80-90 mm Hg to ensure a good early perfusion of the kidney graft following reperfusion. Achieve this partially by adequate volume management and partially by the administration of norepinephrine (0.1 - 1.0 µg/kg/min as a continuous infusion using the mean arterial blood pressure and heart rate for monitoring the efficiency).
11. Perform end-to-end anastomosis of the renal vein:
    1. After placing two corner stitches using 5-0 polypropylene, suture the back wall in a continuous fashion.
    2. Tie the cranial corner stitch and tie it together with the thread used for the back wall.
    3. After finishing the back wall, use the cranial corner stitch to suture the front wall in a cranio-caudal direction. Flush the vein with aheparinized saline solution (100 I.U./mL). Tie the caudal corner stitch.
       NOTE: In case of a size mismatch between the donor and recipient sides, a small growth-factor can be used to ensure a wide and sufficient anastomosis. There are many possible variations of the porcine renal vein branches. In the case of complex venous anatomy, a modified anastomosis approach is necessary (see Figure 3).
12. Perform the end-to-end anastomosis of the renal artery:
    1. Use a 6-0 polypropylene cranial corner stitch to perform the arterial anastomosis. Placing a further caudal, supporting corner stitch which is later removed, is optional.
    2. Suture the back wall in a continuous fashion using the parachute technique. After arriving at the caudal corner remove the second corner stitch (if applicable).
    3. Suture the front wall with the other end of the double-armed 6-0 polypropylene suture. Flush the artery with a heparinized saline solution (100 I.U./mL). Tie the two threads at the caudal corner.
13. Record the time needed for performing both anastomoses with a target warm ischemia time of <40 min.
14. Reperfuse the kidney by opening the venous vascular clamp and subsequently the arterial clamp. Check for significant bleeding.
15. If no significant bleeding from the anastomoses is observed, unwrap the kidney graft and pour warm normal saline solution in the abdomen covering the reperfused graft.
16. Reposition the graft, if needed, to ensure homogeneous reperfusion and avoid congestion.
17. Administer papaverine topically to the outside of the renal artery and the arterial anastomosis (5 mL undiluted).
18. After reperfusion, infuse 250 mL of 20% glucose solution to induce osmotic dieresis followed by the administration of a single dose of 80 mg of furosemide.
    NOTE: Following this, initial urine production may be observed.
19. To ensure urinary drainage, pass a 12 French pediatric urine catheter through the abdominal wall of the right flank of the animal, retroperitoneally.
20. Secure the catheter in the ureter using ligatures (2-0 polyglactin) and block the catheter with 2 mL saline. Further sutures are used to adapt and secure the ureter to the peritoneum of the abdominal wall (2-0 polypropylene). The catheter is also secured to the skin with at least two single knot sutures (2-0 polypropylene).
21. Close the peritoneal layer over the kidney to prevent dislocation of the kidney graft and kinking of the vascular anastomoses (3-0 polyglactin).
22. Close the abdomen in a similar 4-layer fashion as described earlier for the graft retrieval.
23. Following abdominal closure, maintain normothermia on the OR table.
    NOTE: Mean arterial blood pressure should be maintained over 80 mm Hg until the animal is awake and is in a prone position.
24. Following abdominal closure, use color Doppler ultrasound to ensure adequate arterial and venous perfusion of the kidney graft (Figure 4). Monitor the animal closely until it is fully awake and can walk and drink spontaneously. The animals are given 1 L of Ringer solution during the recovery phase.
25. Subsequently, return the animal to its box in the housing facility.

7. Follow up, sample and data collection

1. Provide the animals water ad libitum as soon as they can drink spontaneously. Provide solid food from postoperative day 1.
2. For postoperative analgesia, administer buprenorphine (0.05 - 0.1 mg/kg) intramuscularly every 8 h for 72 h, give pantozol (40mg i.v.) once a day for 72 h. Provide antibiotic treatment (cefuroxime 35 mg/kg i.v. 2x daily) and thrombosis prophylaxis (500 mg of acetylsalicylic acid from postoperative day 1) during the whole observation period until the end of the experiment.
   NOTE: If bleeding complications occur, aspirin is discontinued.
3. Register continuous telemetry data throughout the observation period. Ensure that the animals are visited at least every 8 h by the veterinary officer and/or by an experienced veterinary technician and their clinical condition is evaluated using a score sheet which is used as a basis to prematurely terminate the experiment if required by the clinical condition of the animal.
   NOTE: These so-called humane endpoint criteria are defined as described previously²⁴.
4. Perform daily sample collection using the central venous line and the percutaneous urinary catheter. Change urinary bags (2,000 mL) 2x daily.
5. Following sample collection or the administration of fluids or drugs, block the central venous catheter with heparinized saline solution (100 I.U./mL) between every use to avoid occlusion and cover it with a new sterile cap.
6. Following the corresponding observation period of 5 to 7 days, sacrifice the animals in deep anesthesia following relaparotomy, sample collection and explantation of the kidney graft. Sacrifice is performed using a single injection of pentobarbital (50 - 60 mg/kg i.v.).
   NOTE: In compliance with the 3R principle, the remaining organs and tissues of the sacrificed animals may be used for various ex vivo research and educational purposes in in-house institutes.

Discussion

The porcine model of KT allows the investigation of novel therapeutic approaches and medical devices in a clinically relevant large animal setting^15,17,21. The anatomical, pathophysiological and surgical-technical similarities between the porcine and human setting can facilitate the clinical interpretation of data and the rapid translation of the findings and techniques into clinical testing^{15,16,17,18,19,21}.

The model of orthotopic kidney auto-transplantation does not only comply with the 3R principle by reducing the numbers of required animals compared to allo-transplantation, e.g. no separate donor animal is required, but also provides a unique opportunity to investigate the effects of IRI and preservation injury without the confounding effects of the immunological response and immunosuppressive drugs^17,21.

Slight modifications of the protocol allow modeling a broad spectrum of clinical situations. To mimic KT using donation after circulatory death (DCD) kidneys, vascular structures are clamped for 30 to 60 min in situ before kidney retrieval, while prolonged cold ischemia times (24 hours and longer) can be applied to model extensive preservation injury^16,17,28,29.

Although, the porcine KT model is surgically less challenging than solid organ transplantation models in small animals (e.g. rats and mice)²⁶, there are multiple technical aspects and pitfalls which have to be kept in mind to improve outcomes and avoid specific complications¹⁷.

Failing to avoid the large lymphatic vessels around the inferior vena cava and the aorta during graft retrieval or implantation due to technical mistake or anatomical variations, can lead to a high output lymphatic fistula and post-operative abdominal fluid collection, infection, and potentially technical failure. Lymphatic vessels should be completely avoided during surgery or closed with 5-0 or 6-0 polypropylene sutures. It is wise to also avoid the use of bipolar or any other coagulation device in case of lymphatic leaks. It usually leads to worsening of the situation. In case of a low output lymphatic leakage, our team has a good experience with the application of fibrin-based collagen patches (e.g., Tachosil)³⁰, however, their high cost limits their application in this setting.

In the present protocol we demonstrate a transperitoneal approach for kidney retrieval and auto-transplantation. This is a major technical difference compared to the clinical situation, where kidney grafts are usually implanted into the iliac fossa using an extraperitoneal approach. Although, most groups use a transperitoneal and an orthotopic approach in the porcine model, heterotopic transplantation to the iliac fossa is also possible in pigs³¹. However, due to the relatively low diameter of the external iliac artery in 30-40 kg pigs and its tendency to vasospasm makes it sometimes difficult to perform the end-to-side anastomosis of the renal artery to the external iliac artery³¹. Concerning the fact that we retrieve the left kidney via a transperitoneal approach to perform a subsequent auto-transplantation, it is more feasible to perform the implantation by reopening the same incision and using a straigtforward orthotopic approach, especially that per-protocol it is also required to remove the native right kidney to ensure that the animal will recover with only one predamaged kindey. The comprehensive description of all possible technical variations of the model is beyond the scope of this protocol and has been summarized by others in comprehensive review articles³¹.

Dislocation of the transplanted kidney graft and consequential kinking of the vascular anastomoses is a major source of failure in the porcine KT model, rapidly leading to vascular occlusion and complete failure of the experiment, due to a surgical complication. To avoid this, following auto-transplantation we close the peritoneal layer over the kidney with a running suture using 3-0 polyglactin. Furthermore, color Doppler ultrasound is performed directly following the implantation of the kidney and abdominal closure, to ensure good arterial and venous perfusion of the kidney graft. Ultrasound is also used daily and on-demand, based on the clinical performance of the animal, to screen for kidney perfusion, post-renal problems (e.g. obstruction or kinking of the urinary catheter), and fluid collection due to lymphatic fistula, bleeding or infection (Figure 4 and Figure 6).

As 24 hours of cold ischemia often leads to functional impairment and delayed graft function, the animals may require on-demand medical therapy if it is considered necessary by the veterinary officer. This may include infusion therapy using 5% glucose and/or Ringer solution administered via the central venous line, furosemide bolus injections (in case of oliguria/anuria depending on the clinical state and laboratory results, 60-80 mg bolus injections up to 200 mg/day), and the oral administration of Sodium Polystyrene Sulfonate (Resonium A) in case of severe hyperkalemia³². To avoid experimental bias, the veterinary officer responsible for the post-transplant veterinary care of the animals must be blinded for the applied treatment and grouping.

Although, the anatomy of the renal artery is rather straightforward in German landrace pigs with usually one artery to reconstruct, there is a wide spectrum of anatomical variations of the renal vein branches which require certain surgical creativity during the venous reconstruction. Frequently two (or more) renal vein branches join on different levels between the kidney hilum and the inferior vena cava. The most frequently observed variations and the possible reconstruction options¹⁷ are shown in Figure 3.

Following the first surgical intervention (day -15, telemetry implantation), all animals receive a porcine jacket which they wear throughout the whole period of the experiments. This provides excellent protection against accidental injuries and dislocation of the implanted catheters and provides room for the storage of the urine collection bags. The use of these jackets is also a feasible solution to eliminate the need for metabolic cages for the assessment of creatinine clearance as a refinement method according to the 3R Principle.

Our housing facility integrates the use of telemetry and video-based peri-operative monitoring. Although, these methods cannot replace the regular visits by the veterinary officer and technicians, they facilitate rapid interventions and improve severity assessment to further refine our experimental settings for the future. There is a wide spectrum of indications for the use of an implantable telemetry device in large animal models³³. Although, close monitoring of clinical paramters following major surgery such as ECG, blood pressure, temperature is considered to be standard in the human clinical setting of a surgical intensive- and intermediate care unit, in experimental surgery monitoring is mostly discontinued when the animal is waking up from anaesthesia^33,34,35. Therefore, telemetry provides a feasible way for the continuous monitoring of these animals. We believe that all these data contribute to the early detection of possible postoperative complication accurately and timely (e.g., haemorrhagic shock, or sepsis detected by increasing temperature, hypotonia and tachycardia). This may facilitate timely intervention (e.g., introduction of therapeutic antibiotic therapy, fluid substitution, discontinuation of anticoagulation, or sacrifice of the animal to avoid suffering). Besides these "real-time" monitoring aspect, our group is currently focusing on the severity assessment and refinement of animal experiments^36,37,38. Retrospective analysis of a large amount of collected telemetry data in these experiments may allow us to better stratify the severity of these kind of surgical interventions and optimize perioperative care (e.g., analgesia) in laboratory animals.

In terms of implantable telemetry, a period of at least 12 days after implantation of the measurement system is recommended to ensure stable and optimal measurement data (based on personal communication). After discussing this issue with various manufacturers providing telemetry solutions for large animals as well as with other research groups using these systems in various experimental settings, we decided to integrate a 14 day period between telemetry implantation and kidney transplantation. During the earlier days, deviations may still occur due to the movement of the animal as the scarring and healing processes are still uncomplete.

Despite its advantages, the above-described model has certain limitations. The complexity and required resources and infrastructure are the most important limitations of the model. The time-consuming experimental protocol, complex techniques, and intense peri-operative follow up necessitate the availability of a significant housing and OR capacity and require the involvement of a larger team, including doctoral fellows, surgeons, veterinary officers, and technicians (Table 1). Therefore, based on our empirical observations, it is usually unfeasible to perform more than two procedures a day. A further disadvantage of the porcine model compared to small animal models is the limited possibility of mechanistic and molecular-biological investigations. In the present protocol only 5-days of follow up was reported. This was suitable to demonstrate the most important experimental characteristics of the model, however, this relatively short follow up may not be sufficient to answer certain specific research question (e.g. long-term recovery of function vs. acute damage). Therefore, a project related extension of the follow up might be necessary. This manuscript describes our current "best-practice" in the experimental setting of porcine orthotopic kidney auto-transplantation. While certain steps are mandatory to successfully establish this model, minor aspects (e.g., the intraoperative use of a bladder catheter, arterial catheter placement to the femoral vs. carotid artery) are facultative and may be avoided/altered at the investigators` discretion. Description and justification of each and every methodical aspect would be beyond the scope of the present protocol and has been discussed elsewhere³¹. Finally, it is also difficult to replicate the exact clinical situation of ECD KT in the porcine model where elderly donors, allografts with acute kidney injury and donors with multiple co-morbidities and chronic diseases such as hypertension, diabetes mellitus or arteriosclerosis represent a major part of the marginal donor pool^8,9.

Notwithstanding the above-mentioned limitations as well as technical and logistical challenges, this well-established and reproducible large animal model of KT provides a unique opportunity to investigate novel therapies and techniques to improve organ preservation and clinical outcomes and represents an excellent platform for younger surgeons to master organ transplantation techniques in a large animal model.

Materials

Name	Company	Catalog Number	Comments
Anesthesia materials, drugs and medications
Aspirin 500mg i.v., powder for solution for injection	Bayer Vital AG, Leverkusen, Germany	4324188	antiplatelet agents
Atropine sulfate solution for injection, 100mg	Dr. Franz Köhler Chemie GmbH, Bensheim, Germany	1821288	parasympatholytic agent, premedication
Bepanthen ointment for eyes and nose	Bayer Vital AG, Leverkusen, Germany	1578675	eye ointment
BD Discardit II syringes, 2ml, 5ml, 10ml,20ml	Becton Dickinson GmbH, Heidelberg, Germany	300928, 309050,309110, 300296	syringes
BD Micolance 3 (20G yellow) Cannula	Becton Dickinson GmbH, Heidelberg, Germany	305888	venous catheter
BD Venflon Pro Safety (20G pink)	Becton Dickinson GmbH, Heidelberg, Germany	4491101	venous catheter
Buprenorphine (Buprenovet)	Bayer Vital AG, Leverkusen, Germany	794-996	analgesia
Cefuroxime 750mg, powder for preparing  injection solution	FRESENIUS KABI Deutschland GmbH, Bad Homburg, Germany	J01DC02	antibiotics
Covidien Hi-Contour, Endotracheal Tube 7,5 with Cuffed Murphy Eye	Covidien Deutschland GmbH,Neustadt/Donau, Germany	COV-107-75E	endotracheal Tube
FENTANYL 0,5 mg Rotexmedica solution for injection	Rotexmedica GmbH Arzneimittelwerk, Trittau, Germany	4993593	opioide analgetic agent
Furosemide-ratiopharm 250 mg/25 ml solution for injection	Ratiopharm GmbH, Ulm, Germany	1479542	loop diuretics
Glucose 5% solution for infusion (500ml, 250ml)	B. Braun Deutschland GmbH & Co. KG, Melsungen, Germany	3705273,03705422	infusion fluid
Glucose 20% solution for infusion	B. Braun Deutschland GmbH & Co. KG, Melsungen, Germany	4164483	osmotic diuresis
Heparin-Sodium 5000 I.E./ml	B. Braun Deutschland GmbH & Co. KG, Melsungen, Germany	15782698	anticoagulant
Isoflurane-Piramal (Isoflurane)	Piramal Critical Care Deutschland GmbH, Hallbergmoos, Germany	9714675	volatile anaesthetic agent
Ketamine (Ketamine hydrochloride) 10%	Medistar Arzneimittelvertrieb GmbH, Ascheberg, Germany	0004230	general anaestetic agent
MIDAZOLAM 15mg/3ml	Rotexmedica GmbH Arzneimittelwerk, Trittau, Germany	828093	hybnotica, sedative agent
NaCl 0,9% solution for infusion (500ml,1000ml)	B. Braun Deutschland GmbH & Co. KG, Melsungen, Germany	864671.8779	infusion fluid
Norepinephrine (Arterenol)	Sanofi-Aventis Deutschland GmbH, Frankfurt, Germany	16180	increase in blood pressure
Organ preservation solution (e.g. HTK)	Dr. Franz Köhler Chemie GmbH, Bensheim, Germany	should be decided based on preference and experimental design	organ preservation
Pantoprazole 40mg/solution for injection	Laboratorios Normon,Madrid, Spain	11068	proton pump inhibitor
Paveron N 25mg/ml solution for injection (Papaverine Hydrochloride)	LINDEN Arzneimittel-Vertrieb-GmbH, Heuchelheim, Germany	2748990	spasmolytic agent for vasodilatation
Pentobarbital (Narcoren)	Boehringer Ingelheim vetmedica GmbH, Ingelheim, Germany	1,204,924,565	used for euthanasia
Propofol 1% (10mg/ml) MCT Fresenius	FRESENIUS KABI Deutschland GmbH, Bad Homburg, Germany	654210	general anaesthetic agent
Ringer solution	B. Braun Deutschland GmbH & Co. KG, Melsungen, Germany	1471411	infusion fluid
Sterofundin ISO solution for infusion (1000ml)	B. Braun Deutschland GmbH & Co. KG, Melsungen, Germany	1078961	Infusion fluid
Stresnil (Azaperone) 40mg/ml	Elanco	797-548	sedative
Urine catheter ruffle 12CH	Wirutec Rüsch Medical Vertriebs GmbH, Sulzbach, Germany	RÜSCH-180605-12	transurethral urinecatheter
Surgical materials
Appose ULC Skin Stapler	Covidien Deutschland GmbH,Neustadt/Donau, Germany	8886803712	skin stapler
Cavafix Certo 375	B. Braun Deutschland GmbH & Co. KG, Melsungen, Germany	4153758	central venous catheter
EMKA Easytel +L-EPTA Transponder	emka TECHNOLOGIES S.A.S,Paris,France	L-EEEETA 100	telemetry transponder
EMKA Reciever and Data Analyzer System	emka TECHNOLOGIES S.A.S,Paris,France	Reviever	telemetry receiver
Feather Disposable Scapel (11)(21)	Feather, Japan	8902305.395	scapel
Prolene 2-0, blue monofil VISI-BLACK, FS needle	Johnson & Johnson Medical GmbH - Ethicon Deutschland, Norderstedt, Germany	EH7038H	skin
Prolene 3-0,blue monofil,FS1 needle	Johnson & Johnson Medical GmbH - Ethicon Deutschland, Norderstedt, Germany	EH7694H	skin
Prolene 5-0 (simply angulated, C1 needle) blue monofil VISI-BLACK	Johnson & Johnson Medical GmbH - Ethicon Deutschland, Norderstedt, Germany	EH7227H	vascular
Prolene 5-0 (double armed, C1 needle) 60cm	Johnson & Johnson Medical GmbH - Ethicon Deutschland, Norderstedt, Germany	KBB5661H	vascular
Prolene 6-0 (double armed, C1 needle) 60cm	Johnson & Johnson Medical GmbH - Ethicon Deutschland, Norderstedt, Germany	EH7228H	vascular
Sempermed derma PF Surgical Gloves Seril Gr. 7, 7.5, 8	Semperit investment Asia Pte Ltd, Singapore	4200782,4200871,4200894	surgical gloves
Sentinex® PRO Surgical Gowns Spunlace XL 150cm	Lohmann & Rauscher GmbH & Co. KG, Neuwied, Germany	19302	surgical gown
Tachosil	Takeda Pharma Vertrieb GmbH & Co. KG, Berlin, Germany	MAXI 9,5 x 4,8 cm	haemostasis
Telasorp Belly wipes (green 45x45cm)	PAUL HARTMANN AG,Heidenheim, Germany	4542437	abdominal towel
Pediatric urine catheter	Uromed Kurt Drews KG, Oststeinbeck, Germany	PZN 03280856	used for the uretero-cutaneus stoma
VICRYL- 0 MH Plus	Johnson & Johnson Medical GmbH - Ethicon Deutschland, Norderstedt, Germany	V324	fascial closure
VICRYL - 3-0, SH1 Plus needle, 75cm	Johnson & Johnson Medical GmbH - Ethicon Deutschland, Norderstedt, Germany	W9114	subcutaneous suture, peritoneal suture,
VICRYL - 3-0, SH1 Plus needle, 4*45cm	Johnson & Johnson Medical GmbH - Ethicon Deutschland, Norderstedt, Germany	V780	subcutaneous suture, peritoneal suture,
VICRYL - ligatures Sutupak purple braided, 3-0	Johnson & Johnson Medical GmbH - Ethicon Deutschland, Norderstedt, Germany	V1215E	threats for ligature
3M™ Standard Surgical Mask 1810F	3M Deutschland GmbH, Neuss, Germany	3M-ID 7000039767	surgical mask
Surgical instruments
Anatomical forceps Standard	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	PZ0260	anatomical forceps
Atraumatic tweezers steel, De Bakey Tip 1,5mm 8"	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	GF0840	anatmical atraumatic forceps
Bipolar forceps 16 cm straight, Branch 0,30 mm pointed, universal fit	Bühler Instrumente Medizintechnik GmbH,Tuttlingen, Germany	08/0016-A	biopolar forceps
Bulldog clamp atraumatic,curved, De bakey 78 mm, 3"	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	GF0900	bulldog clamps
DE BAKEY-SATINSKY vascular clamp 215mm	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	GF1661	vascular clamp
Dissecting scissors Mayo,250 mm, 10"	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	SC2232	Scissors for dissection
Dissecting scissors Metzenbaum-Fino, 260 mm, 101/4"	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	SC2290	Scissors for dissection
Draeger CATO Anesthetic machine with PM8050 Monitor	Dräger, Drägerwerk AG & Co. KGaA, Lübeck, Germany	106782	Ventilation System
Fine Tweezers, ADSON 180 mm	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	ADSONPZ0571	fine forceps
Gosset abdomenal wall spreader	CHIRU-INSTRUMENTE, Kaierstuhl,Germany	09-621512	abdominal retractor
HALSTEAD MOSQUITO,curved, surgical 125mm	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	KL2291	mosquite clamps
HF surgical device ICC 300, Electrocautery	Erbe Elektromedizin Gmbh; Tübingen, Germany	20132-043	cautery, biopolar
MICRO HALSTED-MOSQUITO 100mm, curved	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	KL2187	mosquite clamps
Micro steel needle holder straight 0,5mm, with spring lock	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	MN1324D	microsurgical needle holder
Microsurgical/watermaker tweezers LINZ 150mm 6" Ergo round handle	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	MN0087	fine microsurgical forceps
needle holder Mayo-hegar,190 mm, 71/2"	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	NH1255	needle holder
Overhold Slimline Fig. 0 8 1/2"	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	KL4400	overholds
Sterile Gauze 10X10	Paul HaRTMANN AG,Heidenheim, Germany	401725	sterile gauze
Suction tip OP-Flex Handpiece Yankauer	Pfm Medical AG, Köln, Germany	33032182	suction
surgical forceps Standard 5 3/4"	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	PZ1260	surgical forceps
surgical scissors standard pointed-blunt (thread/cloth scissors)175 mm, 7"	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	SC1522	surgical Scissors
Titanit vascular scissors POTTS-SMITH,185 mm, 71/4"60°	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	SC8562	Pott scissors
Tunneling instrument	Marina Medical Instruments Inc,Davies,US	MM-TUN06025	subcutaneous tunneling
Vessel loops	Medline International Germany GmbH,Kleve, Germany	VLMINB	hold and adjust the vessel
Wound spreaders Weitlander, Stump,110 mm, 41/4"	ASANUS Medizintechnik GmbH, Tuttlingen, Germany	WH5210	wound care
Further material
Heating pad	Eickemeyer - Medizintechnik für Tierärzte KG, Tuttlingen, Germany	648050 MHP-E1220	maintain body temperature during surgery
Laryngoscope, customized	Wittex GmbH, Simbach, Germany	333222230	expose the vocal cord
Rectal temperature probe	Asmuth Medizintechnik, Minden, Germany	ASD-RA4	measure body temperature
Spray wound film	Mepro-Dr. Jaeger und Bergmann GmbH, Vechta, Germany	2830	keep sterile condition
Sterile organ bag	Raguse Gesellschaft für medizinische Produkte, Ascheberg, Germany	800059	organ preservation
swine jacket small, adult Landrasse swine 30-50kg, customized for Emka Telemetry and urinary catheterization	Lomir Biomedical Inc., United Kingdom	SS J1LAPMP	swine jackets to pretect implanted catheters and store urine bag
Ultrasound device, Sonosite Edge-II	FUJIFILM SonoSite GmbH, Frankfurt, Germany	V21822	ultrasound and color Doppler
Urine bag 2000ml Volume	ASID BONZ GmbH, Herrenberg, Germany	2062578	disposable urine bag connected to the uretero-cutaneous fistula catheter