We would like to thank Dr. Arthur Nedder for his help and guidance. This work was supported by the Richard A. and Susan F. Smith President&#39;s Innovation Award, Michael B. Klein and Family, The Sidman Family Foundation, The Michael B. Rukin Charitable Foundation, The Kenneth C. Griffin Charitable Research Fund, and The Boston Investment Council.

All in vivo studies were conducted in accordance with the National Institutes of Health&#39;s guidelines on animal care and use and were approved by the Boston Children&#39;s Hospital&#39;s Animal Care and Use Committee (Protocol 18-06-3715). All animals received humane care in compliance with the Guide for the Care and Use of Laboratory Animals. Figure 1 shows the timeline including anesthesia, surgical preparation, and timepoints for primary outcome measurements of this study.
1. Induction, anesthesia and intubation
<ol>
	<li>To prevent unnecessary stress and discomfort, sedate the swine by intramuscular injection of a mixture of tiletamine/zolazepam 4-6 mg/kg and xylazine 1.1-2.2 mg/kg as well as isoflurane 3% using a face mask.</li>
	<li>Cannulate the ear vein to obtain venous access with a 20 G IV cannula after disinfecting the area with 95% ethanol. Start a maintenance infusion (0.9% NaCl at 5 mL/kg/h).</li>
	<li>Intubate the pig with an endotracheal tube (size 7 for pigs weighing 40-50 kg) once anesthetic adequacy is confirmed. Perform balloon-ventilation with a frequency of 12 breaths/min and transport the pig to the operating theater.</li>
	<li>Place the animal on the operating table in the supine position. Immediately start mechanical positive pressure ventilation with FiO2 0.50, 10 mL/kg tidal volume, and a frequency of 12 breaths/min under continuous capnography.</li>
	<li>Place a pulse oximeter on the ear or the bottom lip for continuous monitoring.</li>
	<li>Maintain normothermia (37 &#176;C) using an air-heated pad.</li>
	<li>To maintain general anesthesia, keep isoflurane administration at 0.5-4% through the endotracheal tube. Throughout the procedure, continuously monitor ECG, arterial blood pressure, temperature, and capnography to measure the depth of anesthesia.</li>
	<li>Insert a Foley catheter to check the fluid status of the animal and monitor the urine output by collecting urine in a drainage bag. 
	&#8203;NOTE: Female swine are preferred over males due to the anatomical features of their urethra which allows easier catheterization.</li>
</ol>
2. Surgical preparation and vascular access
<ol>
	<li>Drape the animal in a sterile fashion.</li>
	<li>Disinfect the right lateral area of the neck by applying betadine and then 95% ethanol for 3 times.</li>
	<li>Perform a cut-down for the catheterization of the right carotid artery and the right jugular vein. Retract the sternocleidomastoid muscle laterally and dissect it down to the right carotid artery and the right jugular vein.</li>
	<li>Insert a 5F angiography sheath in both the artery and the vein. Secure it with a silk 2-0 suture.</li>
	<li>Insert a 5F angiography sheath using the Seldinger technique into the left femoral artery.
	<ol>
		<li>To perform the Seldinger technique puncture the femoral artery using a hollow needle. Insert a soft tip guidewire through the lumen and advance it into the femoral artery.</li>
		<li>Hold the guidewire secure with the hand while removing the needle. Pass the angiography sheath over the guidewire into the femoral artery and withdraw the guidewire. Use ultrasound guidance, if necessary.</li>
	</ol>
	</li>
</ol>
3. Induction of renal ischemia-reperfusion injury
<ol>
	<li>Administer 200 IU/kg sodium heparin intravenously to achieve systemic anticoagulation (target activated clotting time (ACT) &#62; 300 s).</li>
	<li>Perform an angiography by injecting an iodinated contrast agent under fluoroscopy to identify the renal arteries. 
	NOTE: To reduce the risk for contrast-induced nephrotoxicity, dilute the iodinated contrast agent in a 1:1 solution with normal saline. Tabulate the dosage for all animals to ensure equivalent dosing.</li>
	<li>Identify the renal arteries, manually advance the guidewire in the guiding catheter.</li>
	<li>Position the 5F JL4 guiding catheter in the left renal artery through the right carotid artery (Figure 1A).</li>
	<li>Position the second 5F JL4 guiding catheter in the right renal artery through the left femoral artery (Figure 1A).</li>
	<li>Use the guidewires to direct a 5F percutaneous transluminal angioplasty (PTA) dilatation catheter in each renal artery. 
	NOTE: It is preferable to position the balloon at the proximal renal artery so that no branches or collaterals of the renal artery are left patent after balloon inflation.</li>
	<li>Position each balloon catheter in place and connect a pressure line to each catheter.</li>
	<li>Check the presence of arterial pulse waveforms in the pressure monitor to ensure the correct positioning of the catheter.</li>
	<li>Inflate each balloon and aim for a pressure of approximately 2.5 atm inside the balloon (Figure 1B).</li>
	<li>To confirm the cessation of blood flow to the kidneys observe the flattening of the pulse waveform at the tip of the balloon catheter.</li>
	<li>Inject iodinated contrast medium (1:1 dilution) and check for any opacification of the renal vessels. 
	NOTE: It is also possible to fill the balloon with an iodinated contrast agent for visualization of the inflated balloon. However, this method is not as sensitive as pulse waveform flattening to confirm occlusion of the renal arteries.</li>
	<li>After 60 min of occlusion, carefully deflate and remove the balloon catheters from the renal arteries.</li>
	<li>Perform an angiography (using 1:1 diluted contrast medium) to confirm renal artery patency and the establishment of renal reperfusion (Figure 1C).</li>
	<li>Remove the 5F angiography sheath from the left femoral artery.</li>
	<li>Apply firm pressure at the site of catheterization for 30 min.</li>
	<li>Reverse the effect of heparin by the administration of protamine (3 mg/kg) until ACT normalizes.</li>
	<li>To sample urine during the post-operative period, secure a tube to the Foley catheter with a silk 2-0 suture using an interrupted stitch on the skin.</li>
	<li>Leave the angiography sheaths in the right carotid artery and the right jugular vein in place and secure them with a silk 2-0 suture using an interrupted stitch to allow for blood sampling throughout the study.</li>
	<li>Close the neck incision with a silk 2-0 suture using a continuous stitch in 2 layers.</li>
	<li>Administer bupivacaine (3 mg/kg) at the incision site to minimize pain.</li>
	<li>Continue to hydrate the animal with 0.9% NaCl at 5 mL/kg/h for a total of 2 h following the end of ischemia.</li>
	<li>Place a fentanyl patch (25-50 &#181;g/h) on the back of the animal to minimize post-operative pain.</li>
	<li>Administer an intramuscular injection of buprenorphine (0.005-0.1 mg/kg) to minimize post-operative pain.</li>
	<li>Monitor the animal and maintain it on mechanical ventilation until awaking.</li>
</ol>
4. Animal recovery
<ol>
	<li>Following awaking, accommodate the animal in a temperature-controlled room.</li>
	<li>Continue to turn the animal from one lateral side to the other until it regains full consciousness and ability to ambulate.</li>
	<li>Provide water and food ad libitum.</li>
</ol>
5. Functional assessment
<ol>
	<li>Collect the blood and the urine samples according to the desired protocol. 
	NOTE: In this study, the following time points were designated: baseline (1 h following initiation of the hydration protocol and before occlusion of the renal arteries), end of ischemia, and reperfusion (2 h, 6 h, 24 h).</li>
	<li>Collect the arterial and the venous blood samples. Store them in lithium heparin or EDTA coated vacutainers for subsequent analysis. 
	NOTE: Draw blood directly from the catheters in the carotid artery and jugular vein.</li>
	<li>Collect the urine samples from the Foley catheter and store them in 15 mL tubes for analysis. 
	NOTE: Collect the urine from the drainage bag connected to the Foley catheter.</li>
	<li>To determine urine output, empty the drainage bag and collect urine for 1 h. 
	NOTE: For the 6 h timepoint at which a drainage bag is not connected to the Foley catheter, close the tube connected to the Foley catheter for 30 min and then collect the urine with a 60 mL syringe to determine urine output.</li>
</ol>
6. Euthanasia
<ol>
	<li>Following the end of the reperfusion period, perform anesthesia and monitor as described above.</li>
	<li>Continue hydration with 0.9% NaCl at 5 mL/kg/h.</li>
	<li>Use the arterial and the venous catheters for blood sampling and the Foley catheter to determine urine output. Collect the final blood and urine samples and calculate the urine output.</li>
	<li>Perform a 15-cm midline laparotomy incision using a size 10 blade from the xiphoid down to the mid pelvis.</li>
	<li>Use a straight lateral retractor to retract the abdominal skin.</li>
	<li>Dissect the lateral peritoneal attachments of the abdominal wall to expose the right and left retroperitoneum.</li>
	<li>Identify and bluntly dissect both renal arteries and veins.</li>
	<li>Ligate both renal arteries and veins with a 2-0 silk suture and perform bilateral nephrectomies to collect whole tissue specimens for histological and metabolic analysis.</li>
	<li>Euthanize the animal with the preferred method of euthanasia (e.g., exsanguination, pentobarbital)</li>
</ol>

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The authors declare no competing financial interests.

AKI is a common clinical disorder affecting up to 50% of hospitalized adult patients worldwide6,12. A clinically relevant animal model is needed to further investigate the pathophysiology of the disease and potential therapeutic targets. Although there are several murine models replicating AKI, these do not completely mimic their respective clinical scenarios and the anatomy of the human kidney. This study proposes a clinically relevant swine model to allow for t...

Acute kidney injury (AKI) is a commonly diagnosed condition among surgical patients associated with significant morbidity and mortality1,2. Available data show that AKI can affect even half of all hospitalized patients worldwide and leads to 50% mortality rate in patients in the intensive care unit1,3. Despite its high prevalence, current AKI therapy remains limited to preventive strategies, such as fluid management and dialysis. Therefore, there is an ongoing interest in exploring alternative therapies for AKI4,5,6.
AKI is typically classified into pre-renal, intrinsic, and post-renal based on its etiology4,5,6. The majority of surgical patients with AKI are associated with pre-renal causes due to hypovolemia, resulting in ischemia-reperfusion injury (IRI) of the kidneys2. Clinically, urine output decreases, and creatinine levels increase due to decreased renal function. The kidney is a high-metabolic-rate organ and susceptible to ischemia. A highly reproducible large animal model of renal IRI is necessary to obtain a better insight into the pathophysiology of AKI and its potential therapeutic approaches5.
To mimic the clinical scenario of kidney hypoperfusion peri-operatively, a model of bilateral renal artery occlusion is deemed suitable. Previously described models entailing unilateral renal artery occlusion with or without resection of the contralateral kidney do not provide sufficient clinical applicability7,8. Although these models are sufficient for causing AKI, they do not resemble real-life clinical scenarios neither in terms of type nor duration of injury.
The aim of this paper is to present a porcine model of percutaneous bilateral temporary occlusion of the renal arteries by balloon-catheter occlusion under angiography. Bilateral renal artery occlusion mimics the clinical scenario of renal hypoperfusion, followed by the subsequent removal of the balloon for reperfusion9,10. The technical steps are described, including catheterization, catheter guidance, angiography, and hemodynamic monitoring. This method not only allows for a highly controlled and replicable occlusion of the renal arteries, but the percutaneous approach minimizes the impact of the inflammatory response by limiting the amount of insult to the body compared to an open approach.

<table><tbody><tr><td>0.9% sodium chloride injection, usp, 100 ml viaflex plastic container</td><td>Baxter</td><td>2B1302</td><td>For animal hydration</td></tr><tr><td>Agent contrast 100.0ml injection media btl ioversal 74%</td><td>CARDINAL HEALTH</td><td>133311</td><td>For visualizing the vasculature</td></tr><tr><td>Bard Bardia Closed System Urinary Drainage Bag</td><td>BARD Inc</td><td>802001</td><td>For urine collection</td></tr><tr><td>BD Vacutainer K2 EDTA</td><td>BD</td><td>367841</td><td>For blood sample storage</td></tr><tr><td>BD Vacutainer Lithium Heparin</td><td>BD</td><td>366667</td><td>For blood sample storage</td></tr><tr><td>Betadine</td><td>Henry Schein</td><td>6906950</td><td>For skin disinfection</td></tr><tr><td>Bookwalker retractor</td><td>Codman</td><td></td><td>For skin retraction</td></tr><tr><td>Bupivacaine 0.25%</td><td>Hospira</td><td></td><td>Administer at incision site for analgesia</td></tr><tr><td>Buprenorphine SR</td><td>Zoo Pharm</td><td></td><td>10mg/ml bottle, Dose: 0.2mg/kg SC</td></tr><tr><td>Cath angio 5.0 Fr x100.0 cm 0.038 in JR4</td><td>MERIT MEDICAL SYSTEM INC</td><td>7523-21</td><td>For identification of the renal arteries</td></tr><tr><td>Cuffed endotracheal tube</td><td>Emdamed</td><td></td><td>To establish a secure airway for the duration of the operation</td></tr><tr><td>EKG Medtronics- Physiocontrol LifePak 20 Oxygen saturation monitor</td><td>GE Healthcare Madison WI</td><td></td><td>For oxygen saturation monitoring</td></tr><tr><td>Encore 26 inflator</td><td>BOSTON SCIENTIFIC</td><td>710113</td><td>For inflating the balloon catheters</td></tr><tr><td>Ethanol 95% (Ethyl alcohol)</td><td>Henry Schein</td><td></td><td>For skin disinfection</td></tr><tr><td>Fentanyl patch</td><td>Mylan</td><td></td><td>Dose: 25-50mcg/hr, TD</td></tr><tr><td>Gold silicone coated Foley</td><td>TELEFLEX MEDICAL INC</td><td>180730160</td><td>For urine collection</td></tr><tr><td>Heparin sodium</td><td>LEO Pharma A/S</td><td></td><td>Dose: 200 IU/kg IV</td></tr><tr><td>i33 ultrasound machine</td><td>Phillips</td><td></td><td>Use ultrasonographic guidance for femoral catherization if necessary</td></tr><tr><td>Inqwire diagnostic guide wire - 0.035&quot; (0.89 mm) - 260 cm (102&quot;) - 1.5 mm j-tip</td><td>MERIT MEDICAL SYSTEM INC</td><td>6609-33</td><td>For guiding the balloon catheters to the renal arteries</td></tr><tr><td>Intravenous catheter, size 20 gauge</td><td>Santa Cruz Biotechnology</td><td>Inc SC-360097</td><td>For fluid administration</td></tr><tr><td>Isoflurane</td><td>Patterson Veterinary Supply, Inc.</td><td>21283620</td><td>Dose: 3%, INH</td></tr><tr><td>Metzenbaum blunt curved 14.5 cm - 5(3/4)&quot;</td><td>Rudolf Medical</td><td>RU-1311-14M</td><td>For tissue dissection and cutting</td></tr><tr><td>Neonatal disposable transducer kit with 30ml/hr flush device and double 4-way stopcocks for continuous monitoring</td><td>Argon Medical</td><td>041588505A</td><td>For pressure measurement</td></tr><tr><td>Powerflex pro PTA dilatation catheter 6 x 20 mm - shaft length (135cm)</td><td>CARDINAL HEALTH</td><td>4400602X</td><td>For occlusion of the renal arteries</td></tr><tr><td>Pressure monitoring lines mll/mll - 12&quot; clear, mll/mll</td><td>Smiths Medical</td><td>B1571/MX571</td><td>For pressure measurement</td></tr><tr><td>Procedure pack</td><td>Molnlycke Health Care</td><td>97027809</td><td>Surgical drape, gauze pads, syringes, beaker etc</td></tr><tr><td>Protamine</td><td>Henry Schein</td><td>1044148</td><td>For heparin reversal</td></tr><tr><td>Scalpel blade - size #10</td><td>Cardinal Health (Allegiance)</td><td>32295-010</td><td>For the skin incisions</td></tr><tr><td>Stopcock iv 4 way lrg bore rotg male ll adptr strl</td><td>Peoplesoft</td><td>1550</td><td>For connecting tubings</td></tr><tr><td>Straight lateral retractor</td><td>Codman</td><td></td><td>For skin retraction</td></tr><tr><td>Suture perma hnd 18in 2-0 braid silk blk</td><td>CARDINAL HEALTH 1</td><td>A185H</td><td>For suturing incision site and securing catheters</td></tr><tr><td>Syringe contrast injection 10ml fixed male luer red</td><td>MERIT MEDICAL SYSTEM INC</td><td>MSS111-R</td><td>To administer the contrast agent</td></tr><tr><td>Syringe medical 60ml ll plst strl ltx free disp</td><td>CARDINAL HEALTH 1</td><td>BF309653</td><td>For urine collection and flushing of the angiocath</td></tr><tr><td>Tilzolan (tiletamine/zolazepam)</td><td>Patterson Veterinary Supply, Inc.</td><td>07-893-1467</td><td>Dose: 4-6 mg/kg, IM</td></tr><tr><td>Xylazine</td><td>Putney, INC</td><td></td><td>Dose: 1.1-2.2 mg/kg, IM</td></tr></tbody></table>

a large animal model for acute kidney injury by temporary bilateral renal artery occlusion

Acute kidney injury (AKI) is associated with higher risk for morbidity and mortality post-operatively. Ischemia-reperfusion injury (IRI) is the most common cause of AKI. To mimic this clinical scenario, this study presents a highly reproducible large animal model of renal IRI in swine using temporary percutaneous bilateral balloon-catheter occlusion of the renal arteries. The renal arteries are occluded for 60 min by introducing the balloon-catheters through the femoral and carotid artery and advancing them into the proximal portion of the arteries. Iodinated contrast is injected in the aorta to assess any opacification of the kidney vessels and confirm the success of the artery occlusion. This is furtherly confirmed by the flattening of the pulse waveform at the tip of the balloon catheters. The balloons are deflated and removed after 60 min of bilateral renal artery occlusion, and the animals are allowed to recover for 24 h. At the end of the study, plasma creatinine and blood urea nitrogen significantly increase, while eGFR and urine output significantly decrease. The need for iodinated contrast is minimal and does not affect renal function. Bilateral renal artery occlusion better mimics the clinical scenario of perioperative renal hypoperfusion, and the percutaneous approach minimizes the impact of the inflammatory response and the risk of infection seen with an open approach, such as a laparotomy. The ability to create and reproduce this clinically relevant swine model eases the clinical translation to humans.

Functional analysis 
The representative results of this study arise from 6 animals and the data shown are mean &#177; standard error of the mean.Renal function is assessed by determining the urine output, estimated glomerular filtration rate (eGFR), plasma creatine, and blood urea nitrogen (BUN). The biomarkers of renal function are assessed using a portable chemistry analyzer. eGFR is calculated according to the following formula: eGFR =1.879 &#215; BW1.092/PCr0.6 (...

Watch this Scientific Journal Video about A Large Animal Model for Acute Kidney Injury by Temporary Bilateral Renal Artery Occlusion at JoVE.com

A Large Animal Model for Acute Kidney Injury by Temporary Bilateral Renal Artery Occlusion

This study presents a highly reproducible large animal model of renal ischemia-reperfusion injury in swine using temporary percutaneous bilateral balloon-catheter occlusion of the renal arteries for 60 min and reperfusion for 24 h.

Acute kidney injury (AKI) is associated with higher risk for morbidity and mortality post-operatively. Ischemia-reperfusion injury (IRI) is the most ...

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4.4K Views.  Harvard Medical School. This study presents a highly reproducible large animal model of renal ischemia-reperfusion injury in swine using temporary percutaneous bilateral balloon-catheter occlusion of the renal arteries for 60 min and reperfusion for 24 h.

Video: A Large Animal Model for Acute Kidney Injury by Temporary Bilateral Renal Artery Occlusion

Use of a Hanging-weight System for Isolated Renal Artery Occlusion

In hospitalized patients, over 50% of cases of acute kidney injury (AKI) are caused by renal ischemia 1-3. A recent study of hospitalized patients revealed that only a mild increase in serum creatinine levels (0.3 to 0.4 mg/dl) is associated with a 70% greater risk of death than in persons without any increase 1. Along these lines, surgical procedures requiring cross-clamping of the aorta and renal vessels are associated with a renal failure rates of up to 30% 4. Similarly, AKI after cardiac surgery occurs in over 10% of patients under normal circumstances and is associated with dramatic increases in mortality. AKI are also common complications after liver transplantation. At least 8-17% of patients end up requiring renal replacement therapy 5. Moreover, delayed graft function due to tubule cell injury during kidney transplantation is frequently related to ischemia-associated AKI 6. Moreover, AKI occurs in approximately 20% of patients suffering from sepsis 6.The occurrence of AKI is associated with dramatic increases of morbidity and mortality 1. Therapeutic approaches are very limited and the majority of interventional trials in AKI have failed in humans. Therefore, additional therapeutic modalities to prevent renal injury from ischemia are urgently needed 3, 7-9. 
To elucidate mechanisms of renal injury due to ischemia and possible therapeutic strategies murine models are intensively required 7-13. Mouse models provide the possibility of utilizing different genetic models including gene-targeted mice and tissue specific gene-targeted mice (cre-flox system). However, murine renal ischemia is technically challenging and experimental details significantly influence results. We performed a systematic evaluation of a novel model for isolated renal artery occlusion in mice, which specifically avoids the use of clamping or suturing the renal pedicle 14. This model requires a nephrectomy of the right kidney since ischemia can be only performed in one kidney due to the experimental setting. In fact, by using a hanging-weight system, the renal artery is only instrumented once throughout the surgical procedure. In addition, no venous or urethral obstruction occurs with this technique. We could demonstrate time-dose-dependent and highly reproducible renal injury with ischemia by measuring serum creatinine. Moreover, when comparing this new model with conventional clamping of the whole pedicle, renal protection by ischemic preconditioning is more profound and more reliable. Therefore his new technique might be useful for other researchers who are working in the field of acute kidney injury.

A precise murine model for acute kidney injury (AKI) due to ischemia is an important tool to investigate acute kidney injury and possibly find therapeutic tools to treat renal injury. The hanging weight system offers a tool for immediate and reliable renal artery occlusion and reperfusion without causing renal congestion.

In hospitalized patients, over 50% of cases of acute kidney injury (AKI) are caused by renal ischemia 1-3. A recent study of hospitalized patients ...

Normothermic Cardiac Arrest and Cardiopulmonary Resuscitation: A Mouse Model of Ischemia-Reperfusion Injury

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused by whole-body hypoperfusion.5,6 Successful reproduction of whole-body hypoperfusion in rodent models has been fraught with 
difficulty.7-9,9,10 Models which employ focal ischemia have repeatedly demonstrated results which do not translate to the clinical setting, and larger 
animal models which allow for whole body hypoperfusion lack access to the full toolset of genetic manipulation possible in the mouse.11,12 However, in recent 
years a mouse model of cardiac arrest and cardiopulmonary resuscitation has emerged which can be adapted to model AKI.13 This model reliably reproduces 
physiologic, functional, anatomic, and histologic outcomes seen in clinical AKI, is rapidly repeatable, and offers all of the significant advantages of a murine surgical 
model, including access to genetic manipulative techniques, low cost relative to large animals, and ease of use. Our group has developed extensive experience 
with use of this model to assess a number of organ-specific outcomes in AKI.14,15

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.

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused ...

Ischemia-reperfusion Model of Acute Kidney Injury and Post Injury Fibrosis in Mice

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often unreported mortality rates that may confound analyses. Bilateral renal pedicle clamping is commonly used to induce IR-AKI, but differences between effective clamp pressures and/or renal responses to ischemia between kidneys often lead to more variable results. In addition, shorter clamp times are known to induce more variable tubular injury, and while mice undergoing bilateral injury with longer clamp times develop more consistent tubular injury, they often die within the first 3 days after injury due to severe renal insufficiency. To improve post-injury survival and obtain more consistent and predictable results, we have developed two models of unilateral ischemia-reperfusion injury followed by contralateral nephrectomy. Both surgeries are performed using a dorsal approach, reducing surgical stress resulting from ventral laparotomy, commonly used for mouse IR-AKI surgeries. For induction of moderate injury BALB/c mice undergo unilateral clamping of the renal pedicle for 26 min and also undergo simultaneous contralateral nephrectomy. Using this approach, 50-60% of mice develop moderate AKI 24 hr after injury but 90-100% of mice survive. To induce more severe AKI, BALB/c mice undergo renal pedicle clamping for 30 min followed by contralateral nephrectomy 8 days after injury. This allows functional assessment of renal recovery after injury with 90-100% survival. Early post-injury tubular damage as well as post injury fibrosis are highly consistent using this model.

We describe models of moderate and severe ischemia-reperfusion-induced kidney injury in which the mice undergo unilateral renal pedicle clamping followed by simultaneous or delayed contralateral nephrectomy, respectively. These models consistently give rise to renal dysfunction and post-injury fibrosis, but injury severity and survival are dependent on mouse background, age and surgical equipment.

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often ...

Renal Ischaemia Reperfusion Injury: A Mouse Model of Injury and Regeneration

Renal ischaemia reperfusion injury (IRI) is a common cause of acute kidney injury (AKI) in patients and occlusion of renal blood flow is unavoidable during renal transplantation. Experimental models that accurately and reproducibly recapitulate renal IRI are crucial in dissecting the pathophysiology of AKI and the development of novel therapeutic agents. Presented here is a mouse model of renal IRI that results in reproducible AKI. This is achieved by a midline laparotomy approach for the surgery with one incision allowing both a right nephrectomy that provides control tissue and clamping of the left renal pedicle to induce ischaemia of the left kidney. By careful monitoring of the clamp position and body temperature during the period of ischaemia this model achieves reproducible functional and structural injury. Mice sacrificed 24 hr&#160;following surgery demonstrate loss of renal function with elevation of the serum or plasma creatinine level as well as structural kidney damage with acute tubular necrosis evident. Renal function improves and the acute tissue injury resolves during the course of 7 days following renal IRI such that this model may be used to study renal regeneration. This model of renal IRI has been utilized to study the molecular and cellular pathophysiology of AKI as well as analysis of the subsequent renal regeneration.

The mouse model of renal ischaemia reperfusion injury described here comprises of a right nephrectomy that provides control tissue and clamping of the left renal pedicle to induce ischaemia that results in acute kidney injury. This model uses a midline laparotomy approach with all steps performed via one incision.

Renal ischaemia reperfusion injury (IRI) is a common cause of acute kidney injury (AKI) in patients and occlusion of renal blood flow is unavoidable ...

Standardized Colon Ascendens Stent Peritonitis in Rats - a Simple, Feasible Animal Model to Induce Septic Acute Kidney Injury

AKI in septic patients is associated with increased mortality and poor outcome despite major efforts to refine the understanding of its pathophysiology. Here, an in vivo model is presented that combines a standardized septic focus to induce AKI and an intensive care (ICU) setup to provide an advanced hemodynamic monitoring and therapy comparable in human sepsis. Sepsis is induced by standardized colon ascendens stent peritonitis (sCASP). AKI is investigated functionally by measurement of blood and urine samples as well as histologically by evaluation of histopathological scores. Furthermore, the advanced hemodynamic monitoring and the possibility of repetitive blood gas sampling enable a differentiated analysis of severity of induced sepsis.
The sCASP method is a standardized, reliable and reproducible method to induce septic AKI. The intensive care setup, continuous hemodynamic and gas exchange monitoring, low mortality rate as well as the opportunity of detailed analyses of kidney function and impairments are advantages of this setup. Therefore, the described method may serve as a new standard for experimental investigations of septic AKI.

Acute kidney injury (AKI) is a severe complication in critically ill patients and is related with an increased mortality. Here, we present a reliable and reproducible in vivo model to mimic AKI under inflammatory conditions that might contribute towards understanding the pathogenesis of septic AKI.

AKI in septic patients is associated with increased mortality and poor outcome despite major efforts to refine the understanding of its pathophysiology. ...

Immunology and Infection

An Effective Mouse Model of Unilateral Renal Ischemia-Reperfusion Injury

Ischemia-reperfusion injury (IRI) is the leading cause of acute renal failure and is a significant contributor to delayed graft function. Animal models are the only available resources that mimic the complexities of the IRI-associated damage encountered in vivo. This paper describes an effective mouse model of unilateral renal IRI that delivers highly reproducible data. Ischemia is induced by occluding the right renal pedicle for 30 min followed by reperfusion. In addition to the surgical procedure, a sequential overview of the expected physiological and histopathological changes following renal IRI will be provided by comparing data from seven different reperfusion times (4 h, 8 h, 16 h, 1 day, 2 days, 4 days, and 7 days). Critical data for planning experiments ahead, such as mean surgical time, average anesthetic consumption, and body weight changes over time, will be shared. This work will help researchers implement a reliable renal IRI model and select the appropriate reperfusion time that aligns with their intended investigative goals.

Renal ischemia-reperfusion injury is associated with high morbidity and mortality in hospitalized patients. Here, we present a simple and effective mouse model of unilateral renal ischemia-reperfusion injury and provide a sequential overview of representative pathological changes observed in the kidney.

Ischemia-reperfusion injury (IRI) is the leading cause of acute renal failure and is a significant contributor to delayed graft function. Animal models ...

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

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.

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

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 ...

Developing Murine Acute Kidney Injury Model Using Bilateral Ischemia-Reperfusion

Bilateral Renal Ischemia-Reperfusion Model for Acute Kidney Injury in Mice

Acute kidney injury (AKI) is defined as a rapid decline in renal function, in which persistent kidney dysfunction gradually progresses to chronic kidney disease (CKD) due to the irreversible loss of nephrons and their maladaptive repair. In recent years, the incidence of AKI has been increasing concerning diverse etiologies, including volume depletion, sepsis, nephrotoxicity, muscle injury, and major trauma, in which ischemia-reperfusion injury (IRI)&#160;accounts for most episodes. Development of the IRI model in mice is induced by surgical clamping of the renal pedicles, which provides powerful and controllable tools for preclinical models of AKI. Importantly, the IRI model is deployed at different stages of the AKI development, especially in the processes of AKI to CKD. Despite the IRI model being widely practiced in many laboratories, a series of variables still influence the results of this model. Here, we describe the procedure of IRI model development to provide a repeatable and reliable method for researchers to explore the underlying pathogenesis in the development of AKI and the progression of AKI to CKD.

The ischemia-reperfusion injury (IRI) model can be used at different stages of acute kidney injury (AKI) development, especially during AKI to chronic kidney disease (CKD) progression. Here, we describe the procedure for IRI model development in mice via a trans-abdominal approach, clamping renal pedicles via a vascular clip to induce ischemic injury.

Acute kidney injury (AKI) is defined as a rapid decline in renal function, in which persistent kidney dysfunction gradually progresses to chronic kidney ...

Standardized 5/6 Nephrectomy in Rats for Renal Insufficiency and Malperfusion

5/6 Nephrectomy Using Sharp Bipolectomy Via Midline Laparotomy in Rats

Chronic kidney disease (CKD) affects over 10% of the global population, amounting to more than 800 million individuals worldwide. The advances in the treatment of CKD have had a significant impact on patient outcomes. While in the past, CKD was often considered a death sentence, with many patients succumbing to the complications of end-stage renal disease, it is now increasingly being managed as a chronic condition with the availability of dialysis and kidney transplantation, as well as new pharmaceutical developments such as SGLT2 inhibitors or nonsteroidal mineralocorticoid receptor antagonists. 
Yet, there continues to be a growing demand for further exploration into the pathophysiological processes and potential therapeutic interventions. Reliable biological models play a crucial role in facilitating this research. Given the multifaceted nature of renal disease, which encompasses not only cell biology but also vascular microanatomy and endocrine signaling, an appropriate model must possess a level of biological complexity that only an animal model can offer, rendering rodents an obvious choice.
This manuscript, therefore, provides an intricate, systematic protocol for surgically reducing renal parenchyma through midline laparotomy and combined total and partial nephrectomy in rats for survival as well as non-survival applications. It emphasizes the critical role of precise surgical techniques in ensuring consistent and reliable outcomes. Prime examples of potential applications for this model include biomolecular and pharmaceutical studies as well as the development of innovative intraoperative imaging modalities, such as hyperspectral imaging, to objectively visualize and differentiate renal malperfusion.

This manuscript introduces a standardized protocol for a 5/6 nephrectomy by sharp bipolectomy via midline laparotomy in a rat model, aiming to induce a state of renal insufficiency via renal parenchyma reduction with a great degree of methodical precision and low risk of technical error.

Chronic kidney disease (CKD) affects over 10% of the global population, amounting to more than 800 million individuals worldwide. The advances in the ...

Kidney Procurement in a Preclinical Large Animal Model

Machine perfusion has evolved as a viable strategy for ex vivo organ assessment, monitoring, treatment, optimization, as well as to prolong preservation times. Large animal models have been paramount for the development and optimization of these technologies. However, in order to ensure graft quality and data reproducibility, standardized and clinically translatable surgical techniques for organ and tissue procurement should be followed. Thus, here, we describe an optimized protocol for kidney procurement in a preclinical swine model. Kidney recovery is performed using mixed breed (Yorkshire cross/mix) pigs. Briefly, following sterile disinfection and draping of the surgical field, a complete midline incision is performed to gain optimal access to both kidneys. The ureter, renal vein, and artery are dissected until their origin from the inferior vena cava and the aorta, respectively. After complete renal dissection, the ureter is tied and cut distally. The donor animal is then fully heparinized with 100 IU per kg/body weight. Next, the renal artery is clamped close to the aorta, and the renal vein is clamped close to the vena cava using a Satinsky vascular clamp. The kidney graft is then resected, and the renal artery is immediately cannulated back table. The kidney will then be flushed with an ice-cold preservation solution and stored on ice until either machine perfusion or transplantation. Finally, the renal artery stump is ligated with a 2-0 silk ligature, and the vena cava is closed with a 6-0 polypropylene suture. This technique recovers kidneys and simulates either a living (single kidney) or deceased (dual kidney) donor setting. The single kidney recovery offers the advantage to perform a subsequent autotransplantation. In the deceased donor model, blood can be collected prior to euthanasia by inserting blood bag needles directly into the aorta, thereby exsanguinating the animal and providing blood for ex vivo machine perfusion.

Here, we present a protocol that details a surgical model for kidney procurement in a preclinical swine model for subsequent machine perfusion or transplantation.

Machine perfusion has evolved as a viable strategy for ex vivo organ assessment, monitoring, treatment, optimization, as well as to prolong preservation ...