This work was supported by NIH R15 1R15DK106666-01A1 (to J. Liu) and NIH RO1 HL071556 (to J.I. Shapiro).

All animal care and experiments were approved by the Marshall University Institutional Animal Care and Use Committee (IACUC) in accordance with the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals. Male C57BL/6 mice (10-12 weeks old) were housed in a pathogen free animal facility in designated rooms equipped with cages that supply purified air under a 12 h light/dark cycle. Food and water were supplied ad libitum.
1. Surgery Preparation
NOTE: The surgical instruments and materials are obtained from different sources that are not specific to surgery operations. Instruments and materials from other sources can also be used for the same purpose. See the Table of Materials for a list of surgical instruments.
<ol>
	<li>Preparation for the surgery
	<ol>
		<li>Place the following within reach of the operating table: warming pad, halogen lamp, oxygen tank and isoflurane tank, isoflurane vaporizer, hair clipper, hair removal cream, scales, sterilized operating set, sterilized surgical linen, cotton swabs, 70% alcohol, polyhydroxydine solution (containing 1% iodine), 0.9% NaCl solution, 1 mL syringes and needles (30 G), 3-0 and 4-0 silk sutures, buprenorphine and penicillin, antibiotic ointment, and lubricant eye ointment.</li>
		<li>Clean a cage and place a heat pad under the bedding for mice housing after surgery.</li>
	</ol>
	</li>
</ol>
2. Step-one Surgery: Pole Ligation of Left Kidney
NOTE: Maintain sterile conditions during the surgery. Cover the autoclaved surgical instruments with a sterile pad. Keep more than one set of surgical instruments in hand for more than one surgery to prevent cross contamination during the surgery. If the instruments need to be used again, in the case of multiple surgeries, disinfect the instruments with betadine solution and 70% ethanol, and then sterilize in a germinator glass bead sterilizer for 5 min. Disinfect the operating area with 70% ethanol. Wear a gown, mask (to cover nose and mouth), cap (to cover head), and a pair of sterile gloves. Change gloves after each surgery.
<ol>
	<li>Wash and disinfect hands and wear sterile surgical gloves. Wear a head cap and a mask.</li>
	<li>Disinfect the table with 70% alcohol. Apply lubricant eye ointment to both eyes of the mouse to prevent eye dryness while under anesthesia.</li>
	<li>Place the mouse in an induction chamber of an isoflurane vaporizer system and induce anesthesia with a mixture of 2.5% isoflurane with 100% oxygen (0.8 - 1.2 L/min) for 1 - 2 min. 
	NOTE: The efficacy of anesthesia was determined by the absence of pain reflex elicited by pinching the tail, as well as by the absence of palpebral reflex by touching the eyelid.</li>
	<li>Inject penicillin (40,000 unit/kg body weight, IM) and buprenorphine (0.02 mg/kg body weight, SC) to prevent infection and pain during the surgery.</li>
	<li>Place the mouse on a warming pad (initially set at 42 &#176;C) with a rectal thermometer to maintain and monitor the body temperature around 37 &#176;C during the procedure. Place the mouse nose into the nose cone and ventilate with a mixture of 0.8-1.2% isoflurane with 100% oxygen (0.8-1.2 L/min) during the surgery.</li>
	<li>Place the mouse on the right side. Shave the hair of surgical area with a hair clipper and then remove with the hair removal cream. Disinfect the site with polyhydroxydine solution (containing 1% iodine) followed by 70% ethanol. Align autoclaved surgical linen on the surgical area.</li>
	<li>Make a ~1.0 cm lumbar incision and separate the muscle and fascia to expose the left kidney using the scalpel and forceps (Figure 1a).</li>
	<li>Pull out the left kidney very gently by pulling on the perirenal fat with small blunt forceps. In the process, in order not to disturb the adrenal gland, move the adrenal gland upwards from the kidney with forceps.</li>
	<li>Using blunt forceps, gently tease the connective tissue and adrenal gland from the superior end of the kidney towards the middle pylorus of the kidney where blood vessels enter and exit. Identify and isolate the ureter. 
	NOTE: The ureter lies around the inferior half of the kidney embedded in the connective tissue and clearly visible under light.</li>
	<li>Gently grab the connective tissue and separate from the kidney tissue to ensure that the ureter is not ligated.</li>
	<li>Before polar ligations, use a sterile ruler to identify the superior and inferior 1/3 part of kidney. The mouse kidney usually is around 1.25 - 1.5 cm long.</li>
	<li>Ligate using a 3-0 silk suture string and with proper force, approximately 0.4 cm from the superior portion towards the middle of pylorus and similarly, 0.4 cm from the inferior portion towards the pylorus (Figure 1a, dotted lines).</li>
	<li>After ligating both poles (Figure 1b), allow a 2-min observation time to make sure that there is no bleeding. 
	NOTE: In the process, it is very important to avoid ligation of the adrenal gland, renal artery, and ureter. If the left ureter is ligated, the mouse will die after removal of the right kidney in the step-two surgery. The ligation force can restrict blood flow to the poles without causing intra-renal bleeding. Within 1 - 2 min after ligation, the poles are discolored due to the limit of blood flow to the poles.</li>
	<li>Push the left kidney back to the original place. Close the muscle and skin with 4-0 and 3-0 silk suture strings, respectively. Apply antibiotic ointment to the surgical area for better recovery and disinfection.</li>
	<li>After ligation, house the mouse separately in a cage with a heat lamp over the cage so that the bedding area is close to 37 &#176;C (monitored with a thermometer), before placing the animals in individual cages. 
	NOTE: Monitor the mouse until it has regained sufficient consciousness to maintain sternal recumbency.</li>
	<li>Allow easy access to water and food during recovery. Administer penicillin and buprenorphine every 12 h for the first 3 days post-surgery to prevent infection and pain. 
	NOTE: The mouse should recover from the trauma of the surgery within 2 - 3 days and should be back to normal eating, drinking and moving. At this time, house the mouse individually for full recovery. The following must be monitored: drinking, eating, walking patterns, awkward gait, hunched back, ocular and/or nasal discharge, and aggressive behavior. Euthanize the mouse if it shows any sign of seizure, coma, untreatable infection, difficulty to walk, loss of gait, unable to eat and drink, and loss of more than 15% pre-operation weight.</li>
</ol>
3. Step-two Surgery: Removal of Right Kidney
<ol>
	<li>Seven days later, expose the right kidney as described for the left kidney in steps 2.1 to 2.10, except place the mouse on its left side.</li>
	<li>Place the right kidney on the linen and clear surrounding fat and connective tissue with blunt forceps.</li>
	<li>Identify the renal artery and vein, and then place two ligatures (3-0 silk) each with a single loose knot around the vessels. Gently tease the connective tissue and adrenal gland from the superior end of the kidney towards the middle pylorus of the kidney using blunt forceps. Gently displace the ureter from the inferior end and tie all the vessels together (3-0 silk) without causing bleeding.</li>
	<li>Move the two loose ligature knots along the vessels. One towards the abdominal aorta side, and the other one towards the kidney side.</li>
	<li>First tie off the ligature knot (towards the abdominal aorta side) with double knots; a solid knot will change the color of the right kidney. Tie off the ligature knot (towards the kidney side) with double knots. Leave enough space between these two double knots so as to cut the vessels between the two knots without cutting the ligature knots.</li>
	<li>Cut the renal vessels between the two knots and remove the right kidney. Check for possible bleeding of the renal vessels.</li>
	<li>Dry the right kidney using sterile gauze and weigh it.</li>
	<li>Close the muscle incision with 4-0 silk suture string and then close the skin incision with 3-0 silk suture string.</li>
	<li>House the mouse separately in a clean solitary cage with a heat lamp over the cage (step 2.15) with easy access to water and food for at least 24 h to recovery. Administer penicillin and buprenorphine every 12 h for 3 days to prevent infection and pain. 
	NOTE: The mouse should recover from the trauma of the surgery within 2 - 3 days, and should be back to normal eating, drinking and moving.</li>
	<li>After surgery, monitor the mice twice daily for three days, and then once a day for the duration of the experiment. Follow the post-surgical monitoring and treatment procedures described in step 2.16.</li>
</ol>
4. Sham Surgery
<ol>
	<li>For the sham surgery, perform the same surgery procedures, including exposure of the kidney, dissection of tissue, and wound closure, but without the pole ligation of left kidney or removal of right kidney.</li>
</ol>
5. Evaluation of Experimental Uremic Cardiomyopathy
<ol>
	<li>One day before sacrifice, perform a transthoracic echocardiography and capture images18. 
	NOTE: Here, the transthoracic echocardiography was performed with a mouse handling table and rectal thermometer. The echocardiographic images were captured using an 18-38 MHz operating frequency transducer attached to an imaging system. Relative wall thickness (RWT), myocardial performance index (MPI), and left ventricle mass index (LVMI) were calculated as we described18.</li>
	<li>At the time of sacrifice, measure the body weight. Measure hematocrit (HCT) with a HCT centrifuge as per manufacturer&#39;s instructions. Prepare the plasma samples using heparin-coated tubes by centrifuging the blood samples for 10 min at 1,500 x g at 4 &#176;C in a tabletop refrigerated centrifuge. Measure cystatin C, creatinine and BUN using kits following the manufacturer&#39;s instructions. Perform the measurements in duplicate.</li>
	<li>At the end of the experiment, anesthetize the mice with ketamine (90 mg/kg, IP) and xylaxine (10 mg/kg, IP) and sacrifice by thoracotomy. Measure the heart weight after removing it from the chest. Prepare cardiac (LV) and kidney homogenates, as well as determination of Type-1 collagen (COL-1), phosphorylation of c-Src and protein carbonylation, as described before18,19.</li>
</ol>

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The authors have nothing to disclose.

The rat 5/6th PNx model has been widely used to study CKD. Because of the much smaller kidney size in mouse, the classical artery ligation and pole resection are very challenging in mouse models with possible high mortality rates and unexpected bleeding/blood loss.
We adopted a mouse PNx model with pole ligation to overcome the bleeding/blood loss. This PNx model takes less time with improved survival rate and high reproducibility. This pole ligation model develops the phenotypic changes of human uremic cardiomyopathy at the time of four weeks post-surgery, and thus provides a model to study its mechanism(s) and therapeutic target(s).
In this PNx model, the 5/6th PNx is an approximate estimation of renal mass reduction since, and up until now, there is no method to exactly calculate the remnant renal mass of the two kidneys that have different sizes/masses. However, the ligation should be performed in a very similar way to reduce operation variation. We estimate that keeping about 40% of the left kidney (or about 20% of total renal mass) can significantly decrease the mortality rate (by about 20%) without sacrificing the development of the uremic cardiomyopathy phenotype. Another issue is the force of ligation which requires practice.
In comparison with the 5/6th PNx models by pole resection or renal artery ligation15,20, the pole ligation method is easy, quick, and less instrumentally demanding. The possible advantages of this PNx model include the following: 1) avoidance of possible blood bleeding as seen in the PNx model by pole resection or artery ligation, 2) impaired renal function with increase in oxidative stress caused by the inflammation response to the pole ligation, 3) lower mortality rate (about 20%), and 4) quick development of uremic cardiomyopathy phenotype. A successful pole ligation shows a progressive irreversible damage and degradation of the ligated poles. The ligated poles are visible at two weeks post-surgery (Figure 4), and mostly disappear at four weeks post-surgery (Figure 5). Comparison of the ligated poles at two weeks and four weeks post-surgery, indicates the inflammation reaction responding to the pole ligation.
This PNx model may be useful to study the mechanism(s) of drug action and therapeutic drug screening. We recently use this PNx model to investigate the role of the Na/K-ATPase signaling and oxidative stress in the development of uremic cardiomyopathy. Administration of pNaKtide, a specific antagonist of the Na/K-ATPase signaling, not only prevents the development of uremic cardiomyopathy but also reverses the onset of uremic cardiomyopathy18.
The critical steps within the protocol are as follows:1) A more aggressive ligation (remnant left kidney mass is less than 30%) will significantly increase the mortality rate (about 50%). 2) The force of ligation requires practice to reach the point that the ligation restricts the blood supply to the poles without breaking the capsule and kidney tissue. Breaking the kidney tissue leads to intra-renal bleeding that can cause death after removal of the right kidney. A successful ligated pole is discolored within 1 - 2 min after ligation, due to the limit of blood flow to the pole. 3) Special attention is required to avoid ligating the ureter of the left kidney or damaging the ureter when ligating the poles. A mouse with ligated or damaged ureter will not survive after removal of the right kidney. This may be the most noticeable disadvantage of the pole ligation method. 4) Maintenance of body temperature during surgery process and post-surgery care. 5) Control of possible infection using antibiotics.

CKD, also known as chronic renal failure, is a progressive loss of kidney function over time that eventually develops into permanent kidney failure. CKD, from early stage renal disease states to end-stage renal disease (ESRD), is a great risk factor for cardiovascular disease events and mortality, and progressively develops to the clinical phenotype called &#34;uremic cardiomyopathy&#34;1,2,3. The uremic cardiomyopathy in patients with CKD or ESRD is associated with cardiovascular abnormalities, mainly caused by overload of left ventricular (LV) pressure and/or volume, leading to LV hypertrophy (LVH), LV dilation, and LV systolic dysfunction4,5,6. Cardiac fibrosis is another common pathological process of uremic cardiomyopathy that reduces cardiac compliance resulting in LV diastolic dysfunction. Severe cardiac fibrosis can lead to sudden cardiac death even in those without cardiac symptoms7.
The 5/6th PNx is a commonly used CKD animal model for animal studies involving renal failure, uremic cardiomyopathy and hypertension. PNx is achieved by ablation of 5/6 renal parenchyma. The rat model was initially developed with the two most common protocols employed being surgical resection or infarction. The rat PNx model is an extremely useful model to study uremic cardiomyopathy with substantial elevations in blood pressure, cardiac hypertrophy and impaired diastolic function. Later, mouse PNx models, operated with the similar techniques as the rat model, were developed due to the wide availability and ease of making genetic manipulations in the mouse system.
It is well-documented that systemic oxidant stress is a constant feature of both clinical and experimental uremic cardiomyopathy8,9. Furthermore, oxidant stress contributes to the uremic syndrome10, and plays a critical role in the pathogenesis of the cardiac abnormalities seen in uremic cardiomyopathy11,12,13. To this point, we have demonstrated that the rodent 5/6th PNx model causes physiological, morphological, and biochemical features of uremic cardiomyopathy14,15,16,17. In the mouse PNx model described here, PNx-operated mice developed significant oxidative stress, at least partially mediated by Na/K-ATPase signaling function, which is critical in PNx-mediated experimental uremic cardiomyopathy. Attenuation of the Na/K-ATPase signaling not only reduces oxidative amplification, but also ameliorates the phenotypical changes in PNx-mediated experimental uremic cardiomyopathy18.

<table><tbody><tr><td>Iris Scissors, 11.5 cm, Straight</td><td>World Precision Instruments</td><td>501758</td><td></td></tr><tr><td>WPI Swiss Tweezers #5 11 cm, 0.1x0.06 mm Tips</td><td>World Precision Instruments</td><td>504506</td><td></td></tr><tr><td>Jewelers #5 Forceps, 11cm, Straight, Titanium</td><td>World Precision Instruments</td><td>555227F</td><td></td></tr><tr><td>Iris Forceps, 10cm, Straight, Serrated</td><td>World Precision Instruments</td><td>15914</td><td></td></tr><tr><td>Tweezers #3, 11cm, 0.2x0.4mm Tips</td><td>World Precision Instruments</td><td>501976</td><td></td></tr><tr><td>Medesy Iris 4.5&quot; Curved Scissors, Stainless Steel</td><td>Net23</td><td>3512</td><td>www.Net32.com</td></tr><tr><td>Dr. Slick Iris Scissors; 3.5&quot;</td><td>Avid Max</td><td>220-1-965-IrisScsrs-P&amp;nbsp;</td><td>www.Avid Max.com</td></tr><tr><td>Miltex Iris Scissors 4 1/8&quot; Curved</td><td>4mdmedical</td><td>V95-306</td><td>www.4mdmedical.com</td></tr><tr><td>5&quot; Hemostat clap, curved jaw</td><td>PJTool</td><td>4355</td><td>www.pjtool.com</td></tr><tr><td>sklar Knapp Iris scissors, straight and sharp/blunt 4&quot;</td><td>Medical Device depot</td><td>64-3430</td><td>www.medicaldevicedepot.com</td></tr><tr><td>Kelly Hemostatic Forceps straight 5.5&quot;</td><td>Pilgtimmedical</td><td>FA710-50</td><td>www.pilgtimmedical.com</td></tr><tr><td>C57BL/6 mice</td><td>Hilltop Lab Animals Inc.</td><td></td><td></td></tr><tr><td>&amp;nbsp;Mouse Handling Table and rectal thermometer</td><td>Visualsonics</td><td></td><td></td></tr><tr><td>MicroScan transducer&amp;nbsp;</td><td>Vevo 1100</td><td>MS400</td><td></td></tr><tr><td>Vevo 1100 Imaging System&amp;nbsp;</td><td>FUJIFILM VisualSonics Inc.</td><td></td><td></td></tr><tr><td>Cystatin C ELISA kit and mouse creatinine kit&amp;nbsp;</td><td>Crystal Chem. Inc.</td><td></td><td></td></tr><tr><td>Mouse BUN ELISA kit&amp;nbsp;</td><td>MyBioSource Inc&amp;nbsp;</td><td></td><td></td></tr></tbody></table>

a mouse 5/6th nephrectomy model that induces experimental uremic cardiomyopathy

Chronic kidney disease (CKD) is a great risk factor for cardiovascular disease events and mortality, and progressively develops to the clinical phenotype called "uremic cardiomyopathy". We describe here an experimental CKD mouse model, named 5/6th partial nephrectomy (PNx) with pole ligation, which developed uremic cardiomyopathy at four weeks post-surgery. This PNx model was performed by a two-step surgery. In step-one surgery, both poles of the left kidney were ligated. In step-two surgery, which was performed 7 days after the step-one surgery, the right kidney was removed. For the sham surgery, the same surgery procedures were performed but without pole ligation of the left kidney or removal of the right kidney. The surgical procedures are easier and less time-consuming, compared to other methods. However, the remnant functional renal mass is not as easily controlled as the renal artery ligation. Four weeks after surgery, in comparison with the sham-operated mice, the PNx mice developed impaired renal function, anemia, cardiac hypertrophy, cardiac fibrosis, and decreased heart systolic and diastolic function.

The data indicated that this modified 5/6th PNx model by pole ligation is a simple and effective model to investigate uremic cardiomyopathy. At four weeks post-surgery, this PNx model presents impaired renal function, anemia, cardiac hypertrophy, cardiac fibrosis, and decreased heart systolic and diastolic function. The results are summarized below.
At four weeks post-surgery, the PNx mice developed impaired renal function in addition to cardiac morphological and biochemical changes that are consistent with the phenotype of human uremic cardiomyopathy. These PNx-mediated changes include the following observations as compared to the sham-operated mice. First, as expected, the PNx surgery caused significant impairment of renal function. As shown in Figure 2, the PNx surgery significantly increased plasma levels of Cystatin C (Figure 2a, n = 16), creatinine (Figure 2b, n = 16), BUN (Figure 2c, n = 16), when compared to the sham surgery. Second, there was PNx-stimulated anemia demonstrated by the significantly lower HCT in the PNx mice (Figure 2d, n=8). Third, the PNx-mediated cardiac hypertrophy and diastolic dysfunction were demonstrated by an increased heart weight/body weight ratio (Figure 3a), and transthoracic echocardiography (ECHO) analysis showing the significant increases in posterior wall thickness (PWT), anterior wall thickness (AWT), RWT (Figure 3b), MPI (Figure 3c), and LVMI (Figure 3d). There was no significant change of ejection fraction (EF, %). Fourth, the PNx-mediated cardiac fibrosis was demonstrated by PNx-induced increases in type I collagen expression in the LV homogenates (Figure 4a) and left remnant kidney homogenates (Figure 4b). Fifth, oxidant stress plays an important role in experimental uremic cardiomyopathy. In this PNx model, PNx significantly stimulated direct protein carbonylation modification of multiple proteins (Figure 4c) and c-Src activation (Figure 4d) in LV homogenates. PNx surgery did not cause significant loss of body weight at four weeks post-surgery. However, the PNx model by pole ligation method did not induce hypertension. There was no significant difference between sham and PNx mice in systolic BP, diastolic BP and mean BP (data not shown) measured at both one day before the step-one surgery and one day before sacrifice, which is consistent with the notion that the C57BL/6 mouse strain is not an established hypertensive model20,21. The data also suggest that this PNx model is unlikely a model of acute renal failure. Moreover, the PNx-mediated cardiac changes, but not impaired renal function, were attenuated by blockage of c-Src activation or induction of heme oxygenase-1 (HO-1)18.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/55825/55825fig1.jpg" /> 
Figure 1: Pole ligation of left kidney. The pictures showed left kidney before ligation (left) and after ligation (right). The doted lines in the left picture showed the estimated ligation lines. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55825/55825fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/55825/55825fig2.jpg" /> 
Figure 2: PNx with pole ligation caused impaired renal function and anemia. At four weeks post-surgery, the PNx surgery significantly increases plasma levels of (a) Cystatin C (n = 16), (b) creatinine (n = 16), (c) BUN (n = 16), compared to the sham surgery. PNx also significantly reduced (d) hematocrit (HCT) in PNx mice (n = 8). Data were expressed as Mean &#177; SEM. **, p &#60;0.01, sham-operated mice vs. PNx-operated mice. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55825/55825fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/55825/55825fig3.jpg" /> 
Figure 3: PNx with pole ligation stimulated cardiac hypertrophy. At four weeks post-surgery, the PNx surgery significantly increases (a) heart weight/body weight ratio (a, n = 8), (b) relative wall thickness (RWT, n =1 9-21), (c) myocardial performance index (MPI, n = 19-21), and (d) left ventricle mass index (LVMI, n = 19-21). Data were expressed as Mean &#177; SEM. **, p &#60;0.01, sham-operated mice vs. PNx-operated mice. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55825/55825fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/55825/55825fig4.jpg" /> 
Figure 4: PNx with pole ligation stimulated type I collagen expression, protein carbonylation and c-Src activation. At four weeks post-surgery, the PNx stimulated type I collagen expression in the left ventricle (LV) homogenates (a, n=11) and left remnant kidney homogenates (b, n = 7-10). Also in LV homogenates, the PNx stimulated direct protein carbonylation (c, n = 6) and c-Src activation (d, n = 6). Data were expressed as Mean &#177; SEM. **, p &#60;0.01, sham-operated mice vs. PNx-operated mice. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55825/55825fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Mouse 5/6th Nephrectomy Model That Induces Experimental Uremic Cardiomyopathy at JoVE.com

A Mouse 5/6th Nephrectomy Model That Induces Experimental Uremic Cardiomyopathy

This manuscript provides a detailed two-step surgical procedure to perform mouse 5/6th partial nephrectomy (PNx) with pole ligation. Four weeks after surgery, in comparison with sham-operated mice, the PNx mice developed impaired renal function, anemia, cardiac hypertrophy, cardiac fibrosis, and decreased heart systolic and diastolic function.

A Mouse 5/6th Nephrectomy Model That Induces Experimental Uremic Cardiomyopathy

Chronic kidney disease (CKD) is a great risk factor for cardiovascular disease events and mortality, and progressively develops to the clinical phenotype ...

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Biology

20.2K Views.  Marshall University. This manuscript provides a detailed two-step surgical procedure to perform mouse 5/6th partial nephrectomy (PNx) with pole ligation. Four weeks after surgery, in comparison with sham-operated mice, the PNx mice developed impaired renal function, anemia, cardiac hypertrophy, cardiac fibrosis, and decreased heart systolic and diastolic function.

Video: A Mouse 5/6th Nephrectomy Model That Induces Experimental Uremic Cardiomyopathy

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 ...

Medicine

Surgical Techniques for Catheter Placement and 5/6 Nephrectomy in Murine Models of Peritoneal Dialysis

Peritoneal dialysis (PD) is a renal replacement therapy consistent on the administration and posterior recovery of a hyperosmotic fluid in the peritoneal cavity to drain water and toxic metabolites that functionally-insufficient kidneys are not able to eliminate. Unfortunately, this procedure deteriorates the peritoneum. Tissue damage triggers the onset of inflammation to heal the injury. If the injury persists and inflammation becomes chronic, it may lead to fibrosis, which is a common occurrence in many diseases. In PD, chronic inflammation and fibrosis, along with other specific processes related to these ones, lead to ultrafiltration capacity deterioration, which means the failure and subsequent cessation of the technique. Working with human samples provides information about this deterioration but presents technical and ethical limitations to obtain biopsies. Animal models are essential to study this deterioration since they overcome these shortcomings.
A chronic mouse infusion model was developed in 2008, which benefits from the wide range of genetically modified mice, opening up the possibility of studying the mechanisms involved. This model employs a customized device designed for mice, consisting of a catheter attached to an access port that is placed subcutaneously at the back of the animal. This procedure avoids continuous puncture of the peritoneum during long-term experiments, reducing infections and inflammation due to injections. Thanks to this model, peritoneal damage induced by chronic PD fluid exposure has been characterized and modulated. This technique allows the infusion of large volumes of fluids and could be used for the study of other diseases in which inoculation of drugs or other substances over extended periods of time is necessary.
This article shows the method for the surgical placement of the catheter in mice. Moreover, it explains the procedure for a 5/6 nephrectomy to mimic the state of renal insufficiency present in PD patients.

This article shows the method for the surgical placement in mice of an intraperitoneal catheter attached to an access port that is positioned at the back of the animal. Moreover, it explains the procedure for a 5/6 nephrectomy to resemble the uremic state of PD patients.

Peritoneal dialysis (PD) is a renal replacement therapy consistent on the administration and posterior recovery of a hyperosmotic fluid in the peritoneal ...

Engineering

Quantitative Real-Time PCR Evaluation of microRNA Expressions in Mouse Kidney with Unilateral Ureteral Obstruction

MicroRNAs (miRNAs) are single stranded, non-coding RNA molecules that typically regulate gene expression at the post-transcriptional level by binding to partially complementary target sites in the 3&#39; untranslated region (UTR) of messenger RNA (mRNA), which reduces the mRNA&#39;s translation and stability. The miRNA expression profiles in various organs and tissues of mice have been investigated, but standard methods for the purification and quantification of miRNA in mouse kidney have not been available. We have established an effective and reliable method for extracting and evaluating miRNA expression in mouse kidney with renal interstitial fibrosis by quantitative reverse-transcription polymerase chain reaction (qRT-PCR). The protocol required five steps: (1) creation of sham and unilateral ureteral obstruction (UUO) mice; (2) extraction of kidney samples from the UUO mice; (3) extraction of total RNA, which includes miRNA, from the kidney samples; (4) complementary DNA (cDNA) synthesis with reverse transcription from miRNA; and (5) qRT-PCR using the cDNA. Using this protocol, we successfully confirmed that compared to the controls, the expression of miRNA-3070-3p was significantly increased and those of miRNA-7218-5p and miRNA-7219-5p were significantly decreased in the kidneys of a mouse model of renal interstitial fibrosis. This protocol can be used to determine the miRNA expression in the kidneys of mice with UUO.

We describe a method for evaluating the microRNA expression in the kidneys of mice with unilateral ureteral obstruction (UUO) by quantitative reverse-transcription polymerase chain reaction. This protocol is suitable for studying kidney microRNA expression profiles in mice with UUO and in the context of other pathological conditions.

MicroRNAs (miRNAs) are single stranded, non-coding RNA molecules that typically regulate gene expression at the post-transcriptional level by binding to ...

Genetics

5/6th Nephrectomy in Combination with High Salt Diet and Nitric Oxide Synthase Inhibition to Induce Chronic Kidney Disease in the Lewis Rat

Chronic kidney disease (CKD) is a global problem. Slowing CKD progression is a major health priority. Since CKD is characterized by complex derangements of homeostasis, integrative animal models are necessary to study development and progression of CKD. To study development of CKD and novel therapeutic interventions in CKD, we use the 5/6th nephrectomy ablation model, a well known experimental model of progressive renal disease, resembling several aspects of human CKD. The gross reduction in renal mass causes progressive glomerular and tubulo-interstitial injury, loss of remnant nephrons and development of systemic and glomerular hypertension. It is also associated with progressive intrarenal capillary loss, inflammation and glomerulosclerosis. Risk factors for CKD invariably impact on endothelial function. To mimic this, we combine removal of 5/6th of renal mass with nitric oxide (NO) depletion and a high salt diet. After arrival and acclimatization, animals receive a NO synthase inhibitor (NG-nitro-L-Arginine) (L-NNA) supplemented to drinking water (20 mg/L) for a period of 4 weeks, followed by right sided uninephrectomy. One week later, a subtotal nephrectomy (SNX) is performed on the left side. After SNX, animals are allowed to recover for two days followed by LNNA in drinking water (20 mg/L) for a further period of 4 weeks. A high salt diet (6%), supplemented in ground chow (see time line Figure 1), is continued throughout the experiment. Progression of renal failure is followed over time by measuring plasma urea, systolic blood pressure and proteinuria. By six weeks after SNX, renal failure has developed. Renal function is measured using 'gold standard' inulin and para-amino hippuric acid (PAH) clearance technology. This model of CKD is characterized by a reduction in glomerular filtration rate (GFR) and effective renal plasma flow (ERPF), hypertension (systolic blood pressure&gt;150 mmHg), proteinuria (&gt; 50 mg/24 hr) and mild uremia (&gt;10 mM). Histological features include tubulo-interstitial damage reflected by inflammation, tubular atrophy and fibrosis and focal glomerulosclerosis leading to massive reduction of healthy glomeruli within the remnant population (&lt;10%). Follow-up until 12 weeks after SNX shows further progression of CKD.

A two-stage method to establish chronic kidney disease (CKD) in the Lewis rat by surgically removing 5/6th of renal mass is described. Combination of the surgical procedure, NOS-inhibition and a high-salt diet leads to a model resembling human CKD, allowing study of causal mechanisms and development of novel therapeutic interventions.

Chronic kidney disease (CKD) is a global problem. Slowing CKD progression is a major health priority. Since CKD is characterized by complex derangements ...

A Murine Model of Irreversible and Reversible Unilateral Ureteric Obstruction

Obstruction of the kidney may affect native or transplanted kidneys and results in kidney injury and scarring. Presented here is a model of obstructive nephropathy induced by unilateral ureteric obstruction (UUO), which can either be irreversible (UUO) or reversible (R-UUO). In the irreversible UUO model, the ureter may be obstructed for variable periods of time in order to induce increasingly severe renal inflammation and interstitial fibrotic scarring. In the reversible R-UUO model the ureter is obstructed to induce hydronephrosis, tubular dilation and inflammation. After a suitable period of time the ureteric obstruction is then surgically reversed by anastomosis of the severed previously obstructed ureter to the bladder in order to allow complete decompression of the kidney and restoration of urinary flow to the bladder. The irreversible UUO model has been used to investigate various aspects of renal inflammation and scarring including the pathogenesis of disease and the testing of potential anti-inflammatory or anti-fibrotic therapies. The more challenging model of R-UUO has been used by some investigators and does offer significant research potential as it allows the study of inflammatory and immune processes and tissue remodeling in an injured and scarred kidney following the removal of the injurious stimulus. As a result, the R-UUO model offers investigators the opportunity to explore the resolution of kidney inflammation together with key aspects of tissue repair. These experimental models are of relevance to human disease as patients often present with obstruction of the renal tract that requires decompression and are commonly left with significant residual kidney impairment that has no current treatment options and may lead to eventual end stage kidney failure.

The murine model of irreversible unilateral ureteric obstruction (UUO) is presented together with the model of reversible UUO in which the ureteric obstruction is relieved by anastomosis of the severed ureter into the bladder. These models enable the study of renal inflammation and scarring as well as tissue remodeling.

Obstruction of the kidney may affect native or transplanted kidneys and results in kidney injury and scarring. Presented here is a model of obstructive ...

A Modified Two Kidney One Clip Mouse Model of Renin Regulation in Renal Artery Stenosis

Renal artery stenosis is a common condition in patients with coronary or peripheral vascular disease where the renin angiotensin aldosterone system (RAAS) is overactivated. In this context, there is a narrowing of the renal arteries that stimulate an increase in the expression and release of renin, the rate-limiting protease in RAAS. The resulting rise in renin expression is a known driver of renovascular hypertension, frequently associated with kidney injury and end organ damage. Thus, there is a great interest in developing novel treatments for this condition. The molecular and cellular mechanism of renin control in renal artery stenosis is not fully understood and warrants further investigation. To induce renal artery stenosis in mice, a modified 2 kidney 1 clip (2K1C) Goldblatt mouse model was developed. The right kidney was stenosed in wild type mice and sham operated mice were used as control. After renal artery stenosis, we determined renin expression and kidney injury. Kidneys were harvested, and fresh cortices were used to determine protein and mRNA expression of renin. This animal model is reproducible and can be used to study pathophysiological responses, molecular and cellular pathways involved in renovascular hypertension and kidney injury.

A modified 2 kidney 1 clip (2K1C) Goldblatt mouse model was developed using polyurethane tubing to initiate renal artery stenosis, inducing an increase in renin expression and kidney injury. Here, we describe a detailed procedure of preparing and placing the cuff onto the renal artery to generate a reproducible and consistent 2K1C mouse model.

Renal artery stenosis is a common condition in patients with coronary or peripheral vascular disease where the renin angiotensin aldosterone system (RAAS) ...

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 ...

A Murine Model of Hemodialysis Access-Related Hand Dysfunction

Chronic kidney disease is a major public health problem, and the prevalence of end-stage renal disease (ESRD) requiring chronic renal replacement therapies such as hemodialysis continues to increase. Autogenous arteriovenous fistula (AVF) placement remains a primary vascular access option for ESRD patients. Unfortunately, approximately half of the hemodialysis patients experience dialysis access-related hand dysfunction (ARHD), ranging from subtle paresthesia to digital gangrene. Notably, the underlying biologic drivers responsible for ARHD are poorly understood, and no adequate animal model exists to elucidate the mechanisms and/or develop novel therapeutics for the prevention/treatment of ARHD. Herein, we describe a new mouse model in which an AVF is created between the left common iliac artery and vein, thereby facilitating the assessment of limb pathophysiology. The microsurgery includes vessel isolation, longitudinal venotomy, creation of arteriovenous anastomosis, and venous reconstruction. Sham surgeries include all the critical steps except for AVF creation. Iliac AVF placement results in clinically relevant alterations in central hemodynamics, peripheral ischemia, and impairments in hindlimb neuromotor performance. This novel preclinical AVF model provides a useful platform that recapitulates common neuromotor perturbations reported by hemodialysis patients, allowing researchers to investigate the mechanisms of ARHD pathophysiology and test potential therapeutics.

This protocol details the surgical steps of murine common iliac arteriovenous fistula creation. We developed this model to study hemodialysis access-related limb pathophysiology.

Chronic kidney disease is a major public health problem, and the prevalence of end-stage renal disease (ESRD) requiring chronic renal replacement ...

Murine Model for Diabetic Cardiomyopathy via High-Fat Diet and STZ Injection

Modeling and Evaluation of Murine Diabetic Cardiomyopathy Model

The underlying pathophysiological mechanisms of diabetic cardiomyopathy (DbCM), a leading cause of mortality among patients with type 2 diabetes mellitus (T2DM), remain poorly understood. The myocardial toxicity associated with T2DM is attributed to factors such as lipotoxicity, glucotoxicity, oxidative stress, reduced cardiac efficiency, and lipoapoptosis. Compared to rats, mice offer greater accessibility, cost-effectiveness, and broader applicability for animal experiments. Insulin resistance and impaired insulin secretion are crucial factors in the pathophysiology of T2DM. We introduce a novel nongenetic murine model that replicates the progression of human DbCM induced by a combination of high-fat diet (HFD) feeding and streptozotocin (STZ) injection. In this study, we used wild-type C57BL/6J mice, administering an HFD regimen for 12 weeks, followed by intraperitoneal injections of STZ for an additional 12 weeks to induce characteristic manifestations of T2DM. We conducted oral glucose tolerance tests and measured serum insulin concentrations to confirm the development of insulin resistance and insufficient insulin secretion. Cardiac structure and function were rigorously assessed through noninvasive transthoracic echocardiography. Pathological characteristics were evaluated through Masson's trichrome staining and wheat germ agglutinin (WGA) staining, revealing pathological features related to DbCM. Therefore, we provide a robust and versatile method for establishing a nongenetic murine model of DbCM.

This protocol outlines a method for inducing diabetic cardiomyopathy through a combination of high-fat diet feeding and streptozotocin injection. This approach aims to provide a reliable framework for scientific investigation into diabetic cardiomyopathy and to explore potential avenues for clinical treatment applications.

The underlying pathophysiological mechanisms of diabetic cardiomyopathy (DbCM), a leading cause of mortality among patients with type 2 diabetes mellitus ...

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 ...

A Mouse 5/6^th Nephrectomy Model That Induces Experimental Uremic Cardiomyopathy

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