Technical Refinement of a Bilateral Renal Ischemia-Reperfusion Mouse Model for Acute Kidney Injury Research

Summary

This study established a protocol focusing on the technical refinement of a mouse model of bilateral renal ischemia-reperfusion for acute kidney injury research.

Abstract

Cardiac arrest poses a large public health burden. Acute kidney injury (AKI) is an adverse marker in survivors of cardiac arrest following the return of spontaneous circulation (ROSC) after successful cardiopulmonary resuscitation. Conversely, recovery of kidney function from AKI is a predictor of favorable neurological outcomes and hospital discharge. However, an effective intervention to prevent kidney damage caused by cardiac arrest after ROSC is lacking, suggesting that additional therapeutic strategies are required. Renal hypoperfusion and reperfusion are two pathophysiological mechanisms that cause AKI after cardiac arrest. Animal models of ischemia-reperfusion-induced AKI (IR-AKI) of both kidneys are comparable with patients with AKI following ROSC in a clinical setting. However, IR-AKI of both kidneys is technically challenging to analyze because the model is associated with high mortality and wide variation in kidney damage, which may affect the analysis. Lightweight mice were chosen, placed under general anesthesia with isoflurane, subjected to surgery with a dorsolateral approach, and their body temperature maintained during operation, thereby reducing tissue damage and establishing a reproducible acute renal IR-AKI research protocol.

Introduction

Cardiac arrest occurs more than 80,000 times annually in the United States^1,2. The mortality rate of cardiac arrest is extremely high^3,4,5,6. AKI is a major risk factor associated with high mortality and poor neurological outcomes in patients with cardiac arrest after ROSC^{7,8,9,10,11,12,13}. Recovery from AKI is a good predictor of favorable neurological outcomes and discharge from the hospital^14,15,16. However, effective therapies for IR-AKI are still lacking^{15,16,17,18,19}. Additional therapeutic strategies are required to further improve the clinical outcomes of the disease.

IR-AKI with bilateral renal ischemia approach is one of the animal models used for AKI research^{20,21,22,23,24,25,26}. Renal IR-AKI animal models are less complicated than a whole-body IR injury model for the study of AKI in patients with sudden cardiac arrest following ROSC^{6,27,28,29,30}. This implies that consistent results from a renal IR-AKI animal model are easier to achieve because of the presence of fewer confounding factors in experiments. Moreover, renal IR-AKI protocols commonly involve a unilateral or bilateral renal pedicle occlusion. Conditions in experiments on bilateral renal IR-AKI are comparable to clinical conditions for AKI following ROSC in patients with sudden cardiac arrest after successful cardiopulmonary resuscitation. Although the pathological characteristics of the kidneys in both models reflect the pathological characteristics of human renal IR injury^31,32,33, a bilateral renal ischemia approach is more relevant to AKI under human pathological conditions, such as cardiac failure, vasoconstriction, and septic shock³⁵. Bilateral renal IR-AKI animal models are suitable for studies focusing on renal IR injuries in cardiac arrest following ROSC.

Bilateral renal IR-AKI models are associated with technical difficulties, experimental complexity, and long surgery duration^{23,26,32,33,35,36}. To overcome these technical difficulties, the present study established a reliable bilateral IR-AKI research protocol in mice by making some technical modifications. The proposed protocol resulted in fewer surgical complications, less tissue damage, and a lower likelihood of mortality during surgery. Therefore, it can be used to investigate the pathophysiological processes of AKI after ROSC to develop new therapeutic strategies against renal hypoperfusion and reperfusion damage^37,38,39.

Protocol

All animal experiments were conducted in accordance with the Guide for the Care and Use of Laboratory Animals, published by the US National Institutes of Health (NIH publication no. 85-23, revised 1996). The study protocol was approved by and in accordance with the guidelines of the Institutional Animal Care and Use Committee at Fu-Jen Catholic University. See the Table of Materials for details about all materials and instruments used in this protocol.

1. Preparing the mice

1. Select 8-week-old C57BL/6 male mice with a weight of 21-23 g.
2. House and maintain the mice under a 12 h light and dark cycle at a controlled temperature (21 ± 2 °C) with free access to food, standard mouse food pellets, and tap water.

2. Anesthesia

1. Put on a surgical mask and sterile gloves.
2. Put the mice under anesthesia with 2% isoflurane mixed with oxygen at 1 L/min in the induction chamber.
3. Assess the level of anesthesia by pedal reflex.
   ​NOTE: Pedal reflex is a retraction of the hind paw in response to a firm toe pinch. The anesthetization is complete when the pedal reflex disappears.
4. Move and place each mouse in prone position on a surgery platform with an electric blanket to maintain their body temperature once the anesthetization is complete. Stabilize the body temperature before surgery and monitor with rectal temperature probes. Apply ophthalmic ointment to both eyes to prevent dryness.
5. Tape the paws of the mice to the board.
6. Attach a mask to the face of the mice to provide a constant supply of 1% isoflurane and 1 L/min oxygen
7. Assess the level of anesthesia by pedal reflex regularly and adjust the anesthetic delivery accordingly during the surgery.

3. Bilateral renal IR-AKI surgery

1. Touch the back and find the lumbar spine of the mice manually. Move along the spine cephalically and look for costovertebral angles that are below both sides of the last rib of the mice.
2. Apply hair removal lotion to both sides of the costovertebral angle region for approximately 30 s and then remove the fur with saline.
3. Disinfect the shaved skin with three rounds of betadine solution and 75% alcohol using cotton balls.
   NOTE: Maintaining a sterile field for surgery throughout the procedure is key. Apply a surgical drape and use sterile instruments.
4. Use fine-tip forceps to gently lift the skin below the left costovertebral angle, and then use scissors to create a 1 cm oblique dorsolateral incision along the skin tension lines from the lumbar midline on the left flank. Transect the muscle wall of the left flank using scissors to visualize the left kidney.
5. Repeat the aforementioned surgical procedures to visualize the right kidney. Remove the small amounts of blood produced during the procedure with sterile cotton swabs.
6. Push and separate the left kidney carefully from the surrounding tissue with forceps. Identify the renal pedicle after the left kidney is exposed.
   NOTE: Be careful not to injure the adrenal gland and the surrounding blood vessels.
7. Clamp over the left renal pedicle with a microvascular clip for 25 min. Confirm ischemia by a visible change in kidney color from pink to dark red.
8. Cover the clamped kidney with sterile saline wet cotton balls to avoid desiccation during left renal pedicle clamping.
9. Repeat the aforementioned surgical procedures to clamp the right renal pedicle with a microvascular clip for 25 min.
10. Cover the clamped kidney with sterile saline wet cotton balls to avoid desiccation during right renal pedicle clamping.
11. Monitor anesthesia depth and humidity of the sterile saline wet cotton balls periodically.
12. Open the left microvascular clip to start reperfusion of the left kidney. Confirm reperfusion by a visible color change of the left kidney from dark red to pink.
13. Open the right microvascular clip to start reperfusion of the right kidney.
14. After the kidney color change is verified, return the kidney to the abdominal cavity.
15. Close the abdominal cavity and skin with 6-0 absorbable suture materials.
16. Scrub to disinfect the wound with a betadine solution and 75% alcohol using cotton balls.
17. Observe the animal carefully until it begins to move freely and feed.
    NOTE: Pay close attention to the animals until they have regained sufficient consciousness to maintain sternal recumbency.
18. Administer carprofen (5 mg/kg in 0.2 mL, administered subcutaneously) for 2-3 days to prevent postsurgical pain.

Results

The quality of the bilateral renal IR-AKI surgery should be assessed before further microscopic or molecular analysis. During surgery, renal ischemia should be confirmed by seeing whether the kidney has changed color from pink to dark red soon after the renal pedicle is clamped with a microvascular clip (Figure 1). After surgery, kidney damage caused by IR-AKI surgery can be further validated with a few microliters of serum through submandibular blood collection for biochemical analysis wher...

Discussion

The proposed bilateral IR-AKI protocol is suitable for investigating the mechanism of hypoperfusion and reperfusion injury of both kidneys. The protocol suggests that lightweight mice, general anesthesia with isoflurane, a dorsolateral approach to the surgery, and body temperature maintenance during the operation mitigate the associated technical difficulties, shorten the duration of the surgery, and increase the consistency of the procedure for acute bilateral renal IR-AKI research.

Technical...

Materials

Name	Company	Catalog Number	Comments
Absorbable Suture, 6-0	Ethicon	J510G-BX
Betadine solution	Shineteh Istrument
Carprofen	Sigma	PHR1452
Cotton balls	Shineteh Istrument
Graefe Forceps	Fine Science Tools	11051-10
Heating pad	Shineteh Istrument
Isoflurane	Piramal Critical Care Inc.	26675-46-7
Moria Vessel Clamp	Fine Science Tools	18320-11
Olsen-Hegar needle holder	Fine Science Tools	12002 - 12
Saline	Shineteh Istrument
Scalpel blades	Shinva	s2646
Small Animal Anesthesia Machine	Sheng-Cing Instruments Co.	STEP AS-01
Tissue scissors	Fine Science Tools	14072 - 10