Name: Mouse Model of Acute to Chronic Kidney Disease Transition Induced by Renal Ischemia/Reperfusion Injury (Video) | JoVE Journal
Uploaded: 2026-02-10T14:00:00.000+00:00
Duration: 7 min 2 s
Description: Learn a dorsal surgical approach for inducing renal ischemia-reperfusion injury in mice to model the acute to chronic kidney disease transition. This protocol achieves high survival and consistent injury for mechanistic studies.

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5 minutes

This work was supported by NIDDK K01-DK133635; UPMC VMI P3HVB award; Samuel and Emma Winters Foundation Award; UPMC CHP RAC Start-up grant; UPMC CHP Foundation Children&#39;s Trust Grantmakers&#39; award; George M. O&#39;Brien Kidney Resource Alliance Opportunity Pool Program (OKRA OPP) (to TC), NIDDK R01-DK121758; DK134346; R25-DK119180 (to SSL), NIDDK R01-DK125015; R01-DK137410 (to JH), and Carolyn and Mark Snyder and the Snyder Family (to JH and SSL). We thank the UPMC CHP Histology Core Laboratory for assistance with histology and scanned imaging, and the Kansas State University Veterinary Diagnostic Laboratories for serum chemistry support.&#160;

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
<ol>
	<li>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.</li>
	<li>Set up a water-circulating heating pad and position an infrared lamp above the surgical field.</li>
	<li>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.</li>
	<li>Transfer the mouse to the heated surgical pad and maintain anesthesia via a nose cone.</li>
	<li>Administer Ethiqa XR (extended-release buprenorphine; 3.25 mg/kg, s.c.) for postoperative analgesia.</li>
	<li>Place the mouse in the prone position on the surgical pad. Secure the limbs with tape if necessary to minimize movement.</li>
	<li>Shave the dorsal hair around the intended incision site using clippers.</li>
	<li>Apply ophthalmic ointment to the eyes to prevent drying during the procedure.</li>
	<li>Insert a rectal probe, secure it with tape, and connect it to a thermometer for continuous monitoring.</li>
	<li>Disinfect the surgical area by alternating 10% povidone-iodine and 70% ethanol, then cover the area with a sterile drape.</li>
</ol>
2. Unilateral renal IRI
<ol>
	<li>Make a midline dorsal skin incision (3-4 cm) spanning the kidney level.</li>
	<li>Separate the skin from the subcutaneous layers using tissue-separating scissors.</li>
	<li>Identify the kidney location by palpation through the skin.</li>
	<li>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.</li>
	<li>Gently exteriorize the kidney through the dorsal incision. If a thin membrane obstructs access, bluntly dissect it.</li>
	<li>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 &#34;canal&#34; for placement of a vascular clamp.</li>
	<li>Confirm that the core body temperature is warm enough (36.8-37.2 &#176;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.</li>
	<li>Pull the skin to cover the kidney to keep it as close to the core body temperature as possible.</li>
	<li>Maintain core body temperature at 36.8-37.2 &#176;C during ischemia by setting the heat pump at 38-39 &#176;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 &#34;sandwich&#34; 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 &#176;C. Record the core body temperature every minute to ensure it is maintained at 36.8-37.2 &#176;C.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
3. Contralateral nephrectomy
<ol>
	<li>Follow step 1, the animal preparation steps.</li>
	<li>Perform a skin incision as described in steps 2.1 and 2.2.</li>
	<li>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.</li>
	<li>While holding the kidney with non-serrated forceps, bluntly dissect the perihilar fat and connective tissue to expose the renal pedicle.</li>
	<li>Carefully separate the adrenal gland from the kidney, leaving the adrenal gland intact and avoiding injury to its vessels.</li>
	<li>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.</li>
	<li>Close the incision as described in step 2.11.</li>
</ol>
4. Collection of blood and kidney tissue
<ol>
	<li>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.</li>
	<li>Place the mouse in the supine position, then make a midline incision and remove the diaphragm.</li>
	<li>Carefully expose the heart and insert a 23-G needle attached to a 3 mL syringe into the left ventricle.</li>
	<li>Draw blood slowly (600-1,000 &#181;L).</li>
	<li>Remove the needle to prevent hemolysis and transfer the blood into the collection tube.</li>
	<li>Excise the left kidney and section it using a scalpel (blade mounted on a handle) according to the study requirements.</li>
	<li>Perform euthanasia by removing the heart under anesthesia (following institutionally approved protocols).</li>
	<li>Process the sectioned tissues according to study requirements (e.g., fixation, snap-freezing).</li>
</ol>

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<li>Skrypnyk, N. I., Harris, R. C., de Caestecker, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ischemia-reperfusion+model+of+acute+kidney+injury+and+post-injury+fibrosis+in+mice%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Ischemia-reperfusion model of acute kidney injury and post-injury fibrosis in mice.</a> J Vis Exp. (78), e50495(2013).</li>
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<li>Chiba, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Endothelial-derived+miR-17%7E92+promotes+angiogenesis+to+protect+against+renal+ischemia-reperfusion+injury%5BTitle%5D)+AND+%22JASN%22%5BJournal%5D)">Endothelial-derived miR-17~92 promotes angiogenesis to protect against renal ischemia-reperfusion injury.</a> JASN. 32 (3), 552-563 (2021).</li>
<li>Silva Barbosa, A. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Dicarboxylic+acid+dietary+supplementation+protects+against+AKI%5BTitle%5D)+AND+%22JASN%22%5BJournal%5D)">Dicarboxylic acid dietary supplementation protects against AKI.</a> JASN. 35, 135-148 (2024).</li>
<li>Chiba, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Loss+of+long-chain+acyl-CoA+dehydrogenase+protects+against+acute+kidney+injury%5BTitle%5D)+AND+%22JCI+Insight%22%5BJournal%5D)">Loss of long-chain acyl-CoA dehydrogenase protects against acute kidney injury.</a> JCI Insight. 10, (2025).</li>
<li>Chiba, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Retinoic+Acid+signaling+coordinates+macrophage-dependent+injury+and+repair+after+AKI%5BTitle%5D)+AND+%22JASN%22%5BJournal%5D)">Retinoic Acid signaling coordinates macrophage-dependent injury and repair after AKI.</a> JASN. 27, 495-508 (2016).</li>
<li>Webster, A. C., Nagler, E. V., Morton, R. L., Masson, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Chronic+kidney+disease%5BTitle%5D)+AND+%22Lancet%22%5BJournal%5D)">Chronic kidney disease.</a> Lancet. 389, 1238-1252 (2017).</li>
<li>Martínez-Klimova, E., Aparicio-Trejo, O. E., Tapia, E., Pedraza-Chaverri, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Unilateral+ureteral+obstruction+as+a+model+to+investigate+fibrosis-attenuating+treatments%5BTitle%5D)+AND+%22Biomolecules%22%5BJournal%5D)">Unilateral ureteral obstruction as a model to investigate fibrosis-attenuating treatments.</a> Biomolecules. 9 (4), 141(2019).</li>
<li>Chevalier, R. L., Forbes, M. S., Thornhill, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ureteral+obstruction+as+a+model+of+renal+interstitial+fibrosis+and+obstructive+nephropathy%5BTitle%5D)+AND+%22Kidney+Int%22%5BJournal%5D)">Ureteral obstruction as a model of renal interstitial fibrosis and obstructive nephropathy.</a> Kidney Int. 75, 1145-1152 (2009).</li>
<li>Bonventre, J. V., Yang, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cellular+pathophysiology+of+ischemic+acute+kidney+injury%5BTitle%5D)+AND+%22J+Clin+Invest%22%5BJournal%5D)">Cellular pathophysiology of ischemic acute kidney injury.</a> J Clin Invest. 121, 4210-4221 (2011).</li>
<li>Wei, Q., Dong, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mouse+model+of+ischemic+acute+kidney+injury%3A+technical+notes+and+tricks%5BTitle%5D)+AND+%22Am+J+Physiol+Renal+Physiol%22%5BJournal%5D)">Mouse model of ischemic acute kidney injury: technical notes and tricks.</a> Am J Physiol Renal Physiol. 303, F1487-F1494 (2012).</li>
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<li>Scarfe, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Long-term+outcomes+in+mouse+models+of+ischemia-reperfusion-induced+acute+kidney+injury%5BTitle%5D)+AND+%22Am+J+Physiol+Renal+Physiol%22%5BJournal%5D)">Long-term outcomes in mouse models of ischemia-reperfusion-induced acute kidney injury.</a> Am J Physiol Renal Physiol. 317, F1068-F1080 (2019).</li>
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</ol>

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 CKD17,18. Unilateral ureteral obstruction (UUO) is a widely used CKD model, which induces obstructive nephropathy and primarily drives fibrosis via persistent tubular obstruction18,<sup class="xr...

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 patients1,2. 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 research4,5,6. 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 fibrosis7,8. 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 publications12,13,14,15. 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.

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

mouse model of acute to chronic kidney disease transition induced by renal ischemia/reperfusion injury

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

Surgical transient occlusion of the renal pedicle induces renal ischemia&#8211;reperfusion injury (IRI), a widely used model for studying acute kidney injury (AKI) and its progression to chronic kidney disease (CKD) in mice. However,&#160;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&#39;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.

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 ...

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.

Surgical transient occlusion of the renal pedicle induces renal ischemia&#8211;reperfusion injury (IRI), a widely used model for studying acute kidney ...

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Research

JoVE Journal

Medicine

A powerful model for perioperative and critical care related acute kidney injury is presented.  Using whole body hypoperfusion induced by cardiac arrest it is possible to nearly replicate the histologic and functional changes of clinical AKI.

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Learn a dorsal surgical approach for inducing renal ischemia-reperfusion injury in mice to model the acute to chronic kidney disease transition. This protocol achieves high survival and consistent injury for mechanistic studies.

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

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