An Effective Mouse Model of Unilateral Renal Ischemia-Reperfusion Injury

The following protocol describes a survival surgery. Therefore, the highest aseptic and surgical practice is applied. All animal experiments were performed in compliance with institutional guidelines and approved by the Institutional Animal Care and Use Committee. To eliminate gender and strain-based differences in IRI effects, only male C57BL6 mice were used in the study. All animals were matched in age and weight to produce comparable results.

1. Preparation

NOTE: A timeline of the different experimental phases and interventions is shown in Figure 1A.

1. Clean and disinfect the surgical table before each procedure. Prepare and place all required materials (sterilized instruments and cotton swabs, sterile gauze and drapes, pre-diluted anesthetics, heating pad, sterilized vascular clamp, sterile saline solution, and skin antiseptics and suture) on the surgical table (see the Table of Materials).
2. Anesthetize male C57BL6 mice (age range 11-13 weeks) by intraperitoneal injection of ketamine/xylazine (100 mg/kg and 20 mg/kg of body weight, respectively, previously diluted in sterile saline).
   NOTE: Skilled animal handling is essential to minimize stress for the animal, as stress responses can negatively affect the action of anesthetics.
3. After ketamine/xylazine administration, shave the surgical area on the right flank using a razor blade and soap.
   NOTE: Shaving the skin improves wound healing as well as the general outcomes of survival surgeries.
4. Disinfect the skin in the surgical area with 70% alcohol first and then with povidone iodine solution using a cotton swab.
5. After the skin preparation, place the mouse on a heating table in a ventral decubitus position and stabilize the body temperature at 37 °C (monitored through rectal and pad sensor probes).
   NOTE: Kidneys are more easily accessed and surgically exposed when placed in ventral decubitus rather than lateral.
6. While the body temperature is stabilized, apply eye ointment to the eyes of the mouse.
   NOTE: Dissociative anesthetics, such as ketamine, cause the animal's eyes to remain open while anesthetized.

2. Surgery

1. Once the pain reflexes are absent (toe pinching with tweezers), perform an approximately 1 cm dorso-lateral surgical incision on the right flank using a scalpel blade. Start the incision behind the last rib and continue caudally approximately 1 cm parallel to the lumbar midline.
2. Transect the abdominal musculature using scissors to visualize the retroperitoneal space. Remove the small amounts of blood produced during the sectioning of the muscles using sterile cotton swabs.
   NOTE: Because the dorso-lateral approach is used, the retroperitoneum, and not the peritoneal cavity, is accessed with this procedure.
3. Push the right kidney out from the abdominal cavity. Use Graefe forceps to expose the kidney carefully.
   NOTE: Always keep forceps closed to avoid traumatic injury to the kidney when placed on the abdomen and use it only to carefully push and guide the kidney towards the surgical incision and out of it.
4. Slowly expose the right kidney and identify the renal pedicle. Carefully remove the adipose tissue around the pedicle.
5. To induce ischemia, place the vascular clamp over the renal artery and vein present in the renal pedicle, avoiding clamping the adjacent ureter. Use a Halsted-Mosquito hemostat for manipulating the vascular clamp.
   NOTE: Ischemia is confirmed by the visualization of a change in color of the kidney from red-pink to dark purple (Figure 1B).
6. Cover the clamped kidney with sterile gauze soaked in saline to avoid desiccation and leave it for 30 min.
7. Monitor anesthesia depth and humidity of the gauze periodically during this time.
   NOTE: The induction dose of anesthesia is sufficient to provide analgesia until the end of the ischemic event; hence, no additional anesthetic injections are required.
8. Shortly before the end of the ischemia period, remove the gauze and uncover the kidney. Hold the Halsted-Mosquito hemostat, ready for clamp removal.
9. At minute 30, open the vascular clamp with the hemostat and remove it from the renal pedicle to allow reperfusion of the kidney.
   NOTE: Reperfusion is confirmed by the visualization of a change in the color of the kidney from dark purple to red-pink (Figure 1C).
10. Perform the same procedures described above for sham animals without clamping of the renal pedicle.
11. After verification of the kidney color change, return the kidney to the abdominal cavity. Close the abdominal muscles with absorbable suture 5-0 using a cruciate pattern.
    NOTE: A second injection of anesthetics may be required to maintain analgesia during the suturing of the muscles and the skin. Half of the initial dose has proved effective in providing analgesia until the conclusion of the surgery.
12. Close the skin with absorbable suture 5-0 using a horizontal mattress pattern. Clean the wound with a povidone iodine solution using a cotton swab.

3. Recovery and post-surgery

NOTE: As post-surgical time is the actual reperfusion time, proper post-surgical care is ethically mandatory and scientifically relevant. Reperfusion times can be selected as required by the researcher. Reperfusion times of 4 h, 8 h, 16 h, 1 day, 2 days, 4 days, and 7 days are compared to obtain a sequential overview of pathological changes induced by renal IRI.

1. Keep the mouse on the heating pad until it starts recovering from anesthesia.
   NOTE: It is recommended to wait until the mouse begins moving its legs and attempts to move around. In cases when additional anesthetic injections are required during the surgery, the recovery time is longer. Atipamezole, an alpha-2 receptor antagonist, can be administered at a dose of 0.5 mg/kg of body weight intraperitoneally to reverse the xylazine effects and shorten the recovery phase. For pain management, buprenorphine (0.1 mg/kg of body weight, intraperitoneally) is administered pre-operatively and every 6 hours during the recovery and post-surgical phase. The use of non-steroidal anti-inflammatory drugs is discouraged as several drugs in this family induce nephrotoxicity and can, therefore, alter the results.
2. After recovery from anesthesia, place the mouse back in its cage with free access to water and food.
   NOTE: Mashed food can be provided in a Petri dish as well as material for hiding and playing (e.g., paper sheets, paper towel tubes).
3. Monitor the mouse daily to assess wound healing, food and water intake, body weight, and behavior.
   NOTE: Wound healing status was assessed using the following scale: 1, dry; 2, wet; 3, partially opened; 4, opened. Fast wound healing was documented in this study, with more than 90% of dry wounds after day 2.

4. Euthanasia and sample collection

1. Euthanize the mice with sodium pentobarbital administered intraperitoneally at a dose that is twice the anesthetic dose for mice (100 mg/kg).
2. Collect fluid and tissue samples as required.
   NOTE: Both kidneys, whole blood (for blood cell count), serum (for blood biochemistry), urine, the heart, and the lungs were collected. A few microliters of serum are needed for blood biochemistry analysis (blood urea nitrogen (BUN), creatinine, electrolytes). If required, 24 h before euthanasia, mice can be placed into metabolic cages to collect a higher urine volume that allows the determination of renal function parameters.