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

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.

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).

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 renal ischemia followed by reperfusion to induce kidney injury. Representative results are presented in Figure 2. Serum was analyzed by the Kansas State Veterinary Diagnostic Laboratory for Renal Profile, which includes analysis of blood urea nitrogen (BUN) and creatinine. Masson's trichrome-stained kidney sections were imaged and analyzed by ImageJ for interstitial collagen deposition. For the real-time qPCR analysis of Acta2 and Col1a1, refer to our previous publication16.

Low-magnification images to capture entire kidney sections demonstrated progressive atrophy 28 and 42 days after renal IRI (Figure 2A). At day 28 and 42 after renal IRI, interstitial fibrosis was observed extensively in the renal cortex and medulla on Masson's trichrome stained kidney tissues (Figure 2B). Serum BUN and creatinine levels are also elevated throughout the course of CKD progression (Figure 2C). Consistent with the histological changes, the expression of widely recognized CKD marker genes (Acta2 and Col1a1) was elevated at days 28 and 42 after renal IRI (Figure 2D).

Representative results obtained using this protocol demonstrate the characteristic results observed when the surgical procedure is performed consistently. Specifically, consistent induction of renal ischemia, careful body temperature management during ischemia, and uniform reperfusion result in reproducible increases in serum renal function markers and elevated expression of genes associated with CKD.

Figure 1: Schematic of the ischemic acute kidney injury to chronic kidney disease (AKI-CKD) mouse model. Please click here to view a larger version of this figure.

Figure 2: Progressive renal interstitial fibrosis after renal IRI. (A) Masson's trichrome-stained kidneys show marked progressive atrophy at days 28 and 42 after renal IRI. Scale bar: 2 mm (All three images are shown at the same magnification). (B) Masson's trichrome-stained kidneys exhibit progressive interstitial collagen deposition in the cortex and medulla (blue area), with tubular atrophy and dilation (*), periglomerular fibrosis (white arrowheads), and perivascular fibrosis (black arrowheads). Scale bars: 50 µm. (C) Increased serum blood urea nitrogen (BUN) and creatinine levels were shown at day 28 and 42 after renal IRI. (D) Fibrosis marker genes (Acta2 and Col1a1) were upregulated at day 28 or 42 after renal IRI. Data are shown as mean ± SD. n=3-6, *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001, using 1-way ANOVA post hoc Tukey multiple comparison. Please click here to view a larger version of this figure.

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^18,19. Renal ischemia-reperfusion induces combined tubular epithelial injury and endothelial/microvascular dysfunction, key mechanisms in ischemic AKI and its progression to CKD^6,20. UUO induces a persistent obstructive insult, whereas our ischemic CKD model is transient. The ischemic CKD model is therefore more appropriate for studies focused on ischemic stress and the AKI-to-CKD transition. In addition, UUO represents a rapidly progressive CKD model, while ischemic CKD develops more slowly and gradually, which may better reflect the course of human CKD. Importantly, this ischemic CKD model permits assessment of renal function, including serum biomarkers and GFR analysis. In this protocol, the most critical factors determining injury consistency and survival rate are strict temperature control during ischemia and maintaining a constant clamping pressure. Specifically, positioning the renal pedicle centrally and vertically within the jaws of the vascular clamp is essential to ensure complete ischemia and to avoid incomplete vascular occlusion, which can lead to variability in injury severity.

Several limitations should be considered when applying it to future studies. Susceptibility to injury is significantly affected by background strain, sex, age, and intraoperative temperature, all of which should be considered in study design²¹. Sensitivity to renal ischemia-reperfusion injury varies among background strains²², and has also been reported to differ even within the same C57BL/6 strain, depending on the vendor²³. While this protocol has been established for FVB/NJ mice, preliminary data confirm that it can also be applied to C57BL/6 mice by using a shorter ischemia duration (approximately 25 min) to induce a CKD phenotype comparable to that observed in FVB/NJ mice. A pilot experiment is essential to establish the optimal duration of renal ischemia for each mouse line of interest that induces a consistent and balanced level of CKD, avoiding both excessive injury and insufficient disease induction. Due to the use of contralateral nephrectomy one day prior to tissue harvest in our renal IRI model, longitudinal assessment at days 28 and 42 cannot be performed. Transdermal GFR measurement can be a useful option to evaluate renal function. It provides a minimally invasive approach for longitudinal assessment in the same animal^24,25. Although the model does not replicate CKD driven by prolonged uremia (as in bilateral renal IRI), it reflects key pathological features of CKD.

Despite these limitations, the protocol can be readily adapted to pharmacologic, dietary, and genetic interventions. It supports diverse mechanistic applications, including cell-type-specific studies, metabolic and mitochondrial modulation, and evaluation of candidate therapies, assessed using prespecified functional, histological, and molecular measures. Its design supports mechanistic studies and the development of novel therapeutic strategies for CKD.