Mouse Model of Acute to Chronic Kidney Disease Transition Induced by Renal Ischemia/Reperfusion Injury

Summary

This article presents an ischemic CKD model using unilateral renal IRI with contralateral nephrectomy performed one day prior to sample collection; this protocol enables serum-based functional assessment, high survival, and reproducible CKD pathology at 28 and 42 days, providing a versatile platform for mechanistic and interventional studies.

Abstract

Surgical transient occlusion of the renal pedicle induces renal ischemia–reperfusion injury (IRI), a widely used model for studying acute kidney injury (AKI) and its progression to chronic kidney disease (CKD) in mice. However, conventional protocols often either preclude serum-based functional assessment or yield high mortality. While existing unilateral and bilateral renal IRI models emphasize reproducibility, challenges remain in achieving both consistent injury severity and long-term survival suitable for studying CKD progression. Here, we present a dorsal approach protocol that provides a robust and reproducible model of the ischemic AKI-to-CKD transition. The dorsal approach facilitates straightforward access to the kidneys and minimizes intra-abdominal manipulation. We use unilateral renal IRI combined with contralateral nephrectomy performed one day prior to blood and tissue collection. This approach offers two main advantages: (1) it minimizes variability by avoiding the inconsistencies associated with bilateral injury, and (2) it achieves nearly 100% survival while maintaining consistent injury, in contrast to protocols in which nephrectomy is performed earlier and associated with higher mortality. Functional evaluation is performed by serum BUN and creatinine, and interstitial fibrosis is quantified by Masson's trichrome-based collagen deposition and CKD marker genes at 28 and 42 days after renal IRI. Together, these readouts provide complementary insight into functional impairment and the development of tissue pathology. This protocol establishes a reliable and versatile model of ischemic-CKD in mice for mechanistic investigations.

Introduction

AKI is a common complication in hospitalized patients and remains a major global health problem. Epidemiological studies indicate that AKI occurs in up to 20% of hospitalized individuals and 30-60% of critically ill patients^1,2. Although some patients recover renal function, others show incomplete recovery, predisposing them to CKD³. The AKI to CKD transition is increasingly recognized as a major determinant of long-term outcomes and has become a focus of mechanistic and therapeutic research^4,5,6. To study the underlying molecular mechanisms and to test potential therapies, reproducible animal models are essential.

Surgical transient occlusion of the renal pedicle (artery and vein) induces renal IRI, one of the most widely used models for studying AKI in mice. This procedure causes acute tubular epithelial injury and an inflammatory response that can progress to chronic interstitial fibrosis^7,8. However, conventional renal IRI protocols present several challenges. Unilateral renal IRI is associated with minimal mortality and is widely employed for long-term investigations⁹; however, assessment of renal function (e.g., serum chemistry, urine analysis, or glomerular filtration rate) is not feasible. Bilateral renal IRI or models combining unilateral renal IRI with simultaneous or early contralateral nephrectomy are associated with increased mortality, limiting their use for chronic studies. While robust variations of these models have been investigated recently¹⁰, their application in chronic study settings warrants careful consideration. In addition, some protocols use a ventral midline laparotomy, which requires manipulation of intra-abdominal organs to expose the renal pedicle. This can increase surgical invasiveness and technical complexity. Skrypnyk et al. reported that contralateral nephrectomy performed 8 days after unilateral ischemia enables serological assessment of renal function after injury, while maintaining a high survival rate¹¹. This protocol provides a systematic set of procedures optimized for assessing the progression from AKI to CKD over a period of up to 42 days.

This study demonstrates an ischemic AKI-CKD model using unilateral renal IRI, followed by delayed contralateral nephrectomy performed one day prior to kidney and blood sample collection. We have previously employed similar procedures as an AKI model in multiple publications^12,13,14,15. We have successfully extended this approach to study the AKI-to-CKD transition, in which, by days 28 and 42, clear signs of CKD are evident (Figure 1 for schematic). This approach provides excellent survival, induces a highly fibrotic phenotype in injured kidneys, and allows assessment of renal function using collected blood. Therefore, this approach is particularly useful for studies requiring serological evaluation (unlike unilateral renal IRI only) and improved survival rate (compared with bilateral renal IRI). A dorsal flank approach was used to access the kidney directly. Compared with ventral midline laparotomy, this approach minimizes intra-abdominal manipulation and confines surgical manipulation to the kidney and perihilar tissues, thereby improving procedural consistency.

Protocol

The animal experiments conducted in this study were approved by the University of Pittsburgh Institutional Animal Care and Use Committee (IACUC; Protocol No. 23103811), in accordance with institutional guidelines. A sham surgery control group can be included as appropriate and typically undergoes surgical exposure without vascular clamping or contralateral nephrectomy. See the Materials section for detailed information on the tools and equipment used. Sharp items, including needles, scalpel blades, and clips, must be disposed of in designated sharps containers. Biological waste, including tissues and cadavers, must be handled and disposed of in accordance with institutional biohazard waste protocols. The reagents and the equipment used are listed in the Table of Materials.

1. Animal preparation and surgical setups

1. Autoclave all surgical instruments prior to use. When performing surgeries on multiple animals, remove blood and debris from the instruments between procedures, then sterilize them for 10-15 s using a hot bead sterilizer. Allow the instruments to cool before reuse.
2. Set up a water-circulating heating pad and position an infrared lamp above the surgical field.
3. Place the mouse in an anesthesia induction chamber and induce anesthesia with isoflurane (1.5%-2% in oxygen, 1-2 L/min) (following institutionally approved protocols. Once the mouse appears to have lost consciousness, confirm adequate anesthetic depth by performing a toe pinch to verify loss of reflexes.
4. Transfer the mouse to the heated surgical pad and maintain anesthesia via a nose cone.
5. Administer Ethiqa XR (extended-release buprenorphine; 3.25 mg/kg, s.c.) for postoperative analgesia.
6. Place the mouse in the prone position on the surgical pad. Secure the limbs with tape if necessary to minimize movement.
7. Shave the dorsal hair around the intended incision site using clippers.
8. Apply ophthalmic ointment to the eyes to prevent drying during the procedure.
9. Insert a rectal probe, secure it with tape, and connect it to a thermometer for continuous monitoring.
10. Disinfect the surgical area by alternating 10% povidone-iodine and 70% ethanol, then cover the area with a sterile drape.

2. Unilateral renal IRI

1. Make a midline dorsal skin incision (3-4 cm) spanning the kidney level.
2. Separate the skin from the subcutaneous layers using tissue-separating scissors.
3. Identify the kidney location by palpation through the skin.
4. At the level where the kidney is visualized, approximately 1 cm lateral to the spine on the left side, make a longitudinal muscle incision of 1-1.5 cm.
   NOTE: If the incision is too long, the kidney may slip back into the abdominal cavity, complicating subsequent manipulation. A minimal skin incision can keep the kidney exteriorized.
5. Gently exteriorize the kidney through the dorsal incision. If a thin membrane obstructs access, bluntly dissect it.
6. While stabilizing the kidney with non-serrated forceps, bluntly dissect the perihilar fat and connective tissue around the upper and lower renal pedicle to create a "canal" for placement of a vascular clamp.
7. Confirm that the core body temperature is warm enough (36.8-37.2 °C) before inducing ischemia. Apply a non-traumatic vascular clamp across the renal artery and vein as a bundle.
   NOTE: Ensure the vessels are centered in the jaws of the clamp and compressed with consistent pressure. Apply the clamp at a consistent location on the hilum. Orient the clamp perpendicular to the vessel axis. This particular clamp can be reused for approximately 10 animal procedures.
8. Pull the skin to cover the kidney to keep it as close to the core body temperature as possible.
9. Maintain core body temperature at 36.8-37.2 °C during ischemia by setting the heat pump at 38-39 °C (adjust as needed) and turning the infrared lamp on/off to maintain the target body temperature (adjust the lamp distance as needed).
   NOTE: This "sandwich" heating system, consisting of a water-circulating heating pad and an infrared lamp, effectively maintains core body temperature within the narrow range of 36.8-37.2 °C. Record the core body temperature every minute to ensure it is maintained at 36.8-37.2 °C.
10. After 30 min of the ischemic period, remove the clamp and return the kidney with a cotton swab to the abdominal cavity.
    NOTE: Confirm the entire kidney appears uniformly pale before declamping. After removing the clamp, confirm successful reperfusion. If reperfusion is markedly delayed, such as 30 s or longer, this may indicate incomplete reperfusion, and exclusion of the animal from subsequent analyses should be considered. Susceptibility to renal IRI and the subsequent severity of CKD are significantly influenced by age, sex, and background strain. Conducting a pilot experiment to determine the optimal ischemia duration is essential for each study design.
11. Close the muscle layer with 5-0 absorbable sutures and then the skin with wound clips (e.g., 9 mm).
    NOTE: Remove wound clips 10 days after surgery.
12. Discontinue isoflurane anesthesia, and allow the mouse to recover in a clean cage. If the animal has recovered from anesthesia and shows no signs of distress or hypothermia, return the cage to the animal maintenance room. Animals are monitored for pain or discomfort post-operatively, and additional doses of Ethiqa XR are administered as required. Humane endpoints are established according to institutional IACUC guidelines to minimize animal distress.
    NOTE: Ethiqa XR provides analgesia for up to 3 days. Administer an additional dose if animals show any signs of pain or discomfort 72 h after the initial dose.

3. Contralateral nephrectomy

1. Follow step 1, the animal preparation steps.
2. Perform a skin incision as described in steps 2.1 and 2.2.
3. At the level where the right kidney is visualized, approximately 1 cm lateral to the spine on the right side, make a longitudinal muscle incision of 1-1.5 cm.
4. While holding the kidney with non-serrated forceps, bluntly dissect the perihilar fat and connective tissue to expose the renal pedicle.
5. Carefully separate the adrenal gland from the kidney, leaving the adrenal gland intact and avoiding injury to its vessels.
6. Ligate renal pedicle with a ligating clip (clip size: small) and 4-0 silk sutures, then transect the vessels on the renal side. Subsequently, transect the ureter and remove the kidney.
7. Close the incision as described in step 2.11.

4. Collection of blood and kidney tissue

1. Induce anesthesia in mice with isoflurane (1.5%-2% in oxygen, 1-2 L/min) and confirm anesthetic depth by toe pinch to verify loss of reflexes.
2. Place the mouse in the supine position, then make a midline incision and remove the diaphragm.
3. Carefully expose the heart and insert a 23-G needle attached to a 3 mL syringe into the left ventricle.
4. Draw blood slowly (600-1,000 µL).
5. Remove the needle to prevent hemolysis and transfer the blood into the collection tube.
6. Excise the left kidney and section it using a scalpel (blade mounted on a handle) according to the study requirements.
7. Perform euthanasia by removing the heart under anesthesia (following institutionally approved protocols).
8. Process the sectioned tissues according to study requirements (e.g., fixation, snap-freezing).

Results

The characteristic change in kidney color observed during surgery confirmed the induction of uniform renal ischemia followed by successful reperfusion. Successful ischemia is indicated by a change in kidney color from black to dark purple, with subsequent improvement after removal of a vascular clamp, confirming reperfusion. If renal ischemia is not successfully induced, the kidney exhibits only minimal color change, and subsequent injury may not be consistent. In this study, FVB/NJ male mice were subjected to 30 min of ...

Discussion

This protocol provides a reproducible renal IRI-induced model of the AKI to CKD transition, which is characterized by sustained kidney dysfunction and progressive interstitial fibrosis, which are widely regarded as hallmarks of CKD^17,18. Unilateral ureteral obstruction (UUO) is a widely used CKD model, which induces obstructive nephropathy and primarily drives fibrosis via persistent tubular obstruction^18,

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Anesthesia gas filter	A.M. Bickford	F-AIR
Anesthesia induction Chamber, warming	AIMS Lab Products	WCS8
Blood collection tube	BD Microtainer	365967
Ethanol	Decon Labs	2701
Ethiqa XR (extended-release buprenorphine)	Fidelis Animal Health	NDC 86084-100-30
Eye ointment	AKORN	NDC 17478-062-35
Gauze pads, sterile	Dynarex	3353
Hair clipper for small animal	WAHL	CLP-9868
Heat lamp, infrared	Physitemp Instruments	HL-1
Heat Pump for warm water circulation	BrainTree Scientific	HTP-1500
Heat/Anesthesia System for small animal, Versaflex	BrainTree Scientific	EZ-7150
Heat-retaining surgical drapes	Braintree Scientific	SPDR-MPS
Hot bead sterilizer	Simon Keller	Steri 250
Isoflurane solution	Covetrus	29405
Ligating clip applier	WECK	533140
Ligating clips	WECK	533837
Microprobe for rectal temperature	Physitemp	RET-3
Microprobe Thermometer	Physitemp	BAT-12
Microscope, optical	Leica	DM2500
Microscope, slider scanner	Leica	Aperio CS2 slide scanner
Mouse (FVB/NJ)	Jackson Laboratory	001800
Needle (23G)	BD	305145
Povidone iodine swabsticks	Dynarex	1201
Scalpel Blades	Fine Science Tools	10020-00
Scalpel Handle	Fine Science Tools	10060-13
Surgical forceps, non-serrated	Roboz Surgical Instrument	RS-5228
Surgical forceps, serrated	Fine Science Tools	11051-10
Surgical forceps, toothed	Fine Science Tools	11019-12
Surgical gloves	CTI	SPGF750
Surgical Tissue separating scissors	Fine Science Tools	14072-10
Suture, Polyglycolic Acid (PGA)/absorbable  (5-0)	Dynarex	9131
Suture, silk (4-0)	Oasis	MV-683-V
Syringe (3ml)	BD	309628
Towel Drapes, sterile	Dynarex	4410
Vascular clamp applying forceps	Fine Science Tools	15057-14
Vascular clamp, non-traumatic	Fine Science Tools	18055-03
Wound clip applier	Fine Science Tools	12020-09
Wound clips	Fine Science Tools	12022-09