The present study was supported by grants from Kidney Research UK (ST4/2011), the Cunningham Trust (CT11/14), the Mrs AE Hogg Charitable Trust for Kidney Research and the Renal Endowment Fund of the Royal Infirmary of Edinburgh.

General Guidance: Animal experiments were performed in accordance with the guidelines and regulations imposed by the Animals (Scientific Procedures) Act 1986. This protocol and the accompanying video protocol are for both a standard UUO and a R-UUO, which can be performed on many mouse strains. In the accompanying video, both procedures are performed on male C57BL/6 mice aged 8 weeks. The data presented in the representative results section were obtained from male FVB/n mice.
NOTE: This protocol and accompanying video details how to perform a standard UUO and R-UUO utilizing the left ureter, however the same techniques can easily be applied to the right ureter.
1. Animal Preparation and Laparotomy
<ol>
	<li>Perform all procedures with sterile (autoclaved) instruments and consumables.</li>
	<li>Inject ketamine hydrochloride (70mg/kg) and medetomidine hydrochloride (1mg/kg) intraperitoneally to anesthetize the mouse. Note: The duration of the resulting anesthetic plane is 4 hr and no supplemental anesthesia is required.</li>
	<li>Confirm the depth of anesthesia by loss of reflexes e.g., toe pinch.</li>
	<li>Remove all hair surrounding the incision area and prepare the abdominal skin for surgery via application of a dilute chlorhexidine solution.</li>
	<li>Place the mouse on a heated surgical pad in a supine position and fix the limbs to the pad using low-tack adhesive tape.</li>
	<li>During the procedure, monitor the mouse for signs of thermal burns as a result of the heated surgical pad. If possible use a non-electric heat source.</li>
	<li>Administer analgesic by a subcutaneous injection of buprenorphine hydrochloride (0.06mg/kg) and apply eye lubricant to prevent corneal drying.</li>
	<li>Make a midline laparotomy and an incision of the avascular linea alba using tissue separating scissors to gain access to the peritoneal cavity.</li>
	<li>Drape the mouse and insert a colibri retractor into the incision.</li>
</ol>
2. Unilateral Ureteric Obstruction
<ol>
	<li>Using sterilized cotton buds expose the left ureter by displacing the intestines towards the right side of the abdominal cavity and cover them with moistened drapes.</li>
	<li>Using angled forceps isolate and lift the left ureter.</li>
	<li>To create a ureteric obstruction, ligate the left ureter twice with 6/O black braided silk suture anywhere between the bladder and renal pelvis. For long term experiments, use absorbable suture for all abdominal surgeries. Alternatively, a ligating clip can be applied to the ureter.</li>
	<li>To isolate the bladder from the ureter, divide the ureter between the two sutures.</li>
	<li>Carefully replace the intestines into the peritoneal cavity.</li>
	<li>Follow the steps listed in Section 4 &#8211; Post-Operative Recovery and Care to close the incision and reverse the anesthesia.</li>
</ol>
3. Preparation for Reversible Unilateral Ureteric Obstruction
<ol>
	<li>Prepare a mouse for surgery and isolate the left ureter as detailed in Steps 1.1 to 2.2 above.</li>
	<li>To create a ureteric obstruction that can be reversed, ligate the left ureter twice with 6/O black braided silk suture close to the bladder. Leave one end of the upper suture long as this will be used to anchor the soft walled plastic tubing in place around the ureter.</li>
	<li>Make a longitudinal slit in a 5mm length of soft walled plastic tubing such that it can be splayed open to allow it to be applied to the ureter. Any soft walled silicone plastic tubing, with an internal diameter of 1mm and external diameter of 2mm, can be used.</li>
	<li>Place the soft walled plastic tubing gently around the ureter. Ensure that the long suture emerges from the center of the slit in the tubing once it is closed around the ureter.</li>
	<li>Place a length of 6/O black braided silk suture around the soft walled tubing enclosing the ureter and tie once. Now place the long end of the suture, emerging from the center of the slit in the tubing, longitudinally across the tubing such that it lies on top of the previously tied suture.</li>
	<li>Tie the suture located around the tubing twice to anchor the tubing and the long suture end in place around the ureter. 
	Note: To prevent adhesion formation on the ureter and the soft walled plastic tubing, an adhesion reduction solution can be applied to the area around the tubing.</li>
	<li>Carefully replace the intestines into the peritoneal cavity.</li>
</ol>
4. Post-Operative Recovery and Care
<ol>
	<li>Close the peritoneum with blanket stitch using 5/O black braided silk suture and approximate the skin using metallic skin clips.</li>
	<li>To minimize the risk of post-operative infection, apply an antiseptic such as iodine/alcohol solution to the abdominal skin.</li>
	<li>Partially reverse anesthesia with atipamezole hydrochloride (2mg/kg) subcutaneously.</li>
	<li>Administer fluids by a subcutaneous injection of 1 ml&#160;warmed saline.</li>
	<li>Monitor the mouse until it has recovered consciousness.</li>
	<li>Allow the mouse to recover in a heated box kept at 29 &#176;C for 24 hr. Moistened food can also be provided to encourage fluid and nutrition intake.</li>
	<li>Leave the mouse to recover to induce a desired level of obstruction, typically 7 days.</li>
	<li>For long-term recovery experiments, provide ongoing analgesics. If the mouse is to be recovered for longer than 7 days, remove the skin clips 7 days following surgery.</li>
	<li>Once the desired level of obstruction has been induced, either reverse the UUO, as described below, or euthanize the mouse by cervical dislocation and collect the kidneys for histopathology analysis.</li>
</ol>
5. Reversible Unilateral Ureteric Obstruction
<ol>
	<li>To perform a R-UUO prepare a mouse, which has undergone preparation for R-UUO, for surgery as described in steps 1.1 to 1.7.</li>
	<li>If present, remove the skin clips and divide or remove the sutures in the peritoneum to gain access to the abdominal cavity. Note: If the ureter has been obstructed for a long-term experiment skin clips should have been removed 7 days following application.</li>
	<li>Prepare the mouse and isolate the left ureter as described in steps 1.8 to 2.2.</li>
	<li>Using angled forceps, free the soft walled plastic tubing from any granulomatous tissue which may have formed.</li>
	<li>Cut the suture holding the plastic tubing around the obstructed ureter with a scalpel and remove the tubing.</li>
	<li>Confirm successful UUO by assessing for the presence of hydronephrosis in the left kidney, the kidney should also appear pale.</li>
	<li>Divide the ureter between the sutures.</li>
	<li>Place the remaining length of ureter, attached to the kidney, on a small piece of sterile gauze. This will be used to collect the urinary sediment and dead cells that will drain from the ureter and renal pelvis once the suture is removed.</li>
	<li>Divide the ureter above, but near, the suture and allow the kidney to drain onto the gauze. Leave the lower suture, closest to the bladder, intact. This is to ensure that no urine will leak from the bladder and into the peritoneal cavity.</li>
	<li>Once the ureter and renal pelvis have been drained, apply a long 6/O black braided silk suture to the end of the remaining length of ureter. This will be used to aid the ureter to bladder anastomosis performed later.</li>
	<li>Follow the steps below to anastomose the remaining length of ureter into the bladder:
	<ol>
		<li>Turn the ureter such that it lies anterior to its original position and lies over the kidney.</li>
		<li>Place a single 9/O polyamide monofilament tacking suture 2 mm from the end of the ureter such that it emerges in the direction of the bladder. Take care to ensure that the suture remains in the muscular coat and does not enter the ureter lumen. The tacking suture will be used to anchor the ureter in the bladder.</li>
		<li>To create a channel through the bladder, pass a 21G needle diagonally through the bladder such that it exits towards the frontal (ventral) wall of the bladder.</li>
		<li>Rest an eyed needle in the bevel of the 21G needle. Use the 21G needle to guide the eyed needle through the bladder. The eyed needle will be used to take the ureter through the bladder.</li>
		<li>Pass the 9/O tacking suture, applied to the ureter in step 5.11.2, through the first incision in the bladder and out through the bladder wall adjacent to the entry point. Once the ureter is passed through the bladder this will be tied to anchor the ureter to the bladder.</li>
		<li>Place the long 6/O black braided silk suture applied to the end of the ureter in step 5.10 through the eyed needle.</li>
		<li>Carefully withdraw the eyed needle out of the bladder whilst ensuring that the ureter is also pulled through the bladder.</li>
		<li>Once the ureter emerges from the bladder remove the eyed needle and apply a clamp to the long suture at the end of the ureter to prevent retraction back into the bladder.</li>
		<li>To anchor the ureter within place in the bladder, tie the 9/O polyamide monofilament tack suture, applied in step 5.11.5.</li>
		<li>Apply additional single 9/O polyamide monofilament tie sutures as described in steps 5.11.2 and 5.11.5 at two locations around the entry point to firmly anchor the ureter within the bladder.</li>
		<li>Using the back of opened scissors, push the bladder wall back slightly to expose more of the emergent ureter.</li>
		<li>Divide the ureter above the long suture located at the end of the ureter. The ureter should retract into the body of the bladder. It is usual to observe urine flowing out of the opening in the bladder, confirming the open lumen of the ureter.</li>
		<li>Close the exit wound in the bladder with a 9/O polyamide monofilament suture.</li>
	</ol>
	</li>
	<li>Carefully replace the intestines into the peritoneal cavity.</li>
	<li>Provide post-operative care as detailed in steps 4.1 to 4.8.</li>
	<li>Allow the mouse to recover until the kidney is decompressed, typically 7 days.</li>
	<li>At the experimental end point, euthanize the mouse by cervical dislocation and collect the kidneys for histopathological analysis.</li>
</ol>

<ol>
	<li>Chevalier, R. L., Forbes, M. S., Thornhill, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ureteral+obstruction+as+a+model+of+renal+interstitial+fibrosis+and+obstructive+nephropathy.">Ureteral obstruction as a model of renal interstitial fibrosis and obstructive nephropathy.</a> Kidney Int. 75, 1145-1152 (2009).</li><li>Kipari, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nitric+oxide+is+an+important+mediator+of+renal+tubular+epithelial+cell+death+in+vitro+and+in+murine+experimental+hydronephrosis.">Nitric oxide is an important mediator of renal tubular epithelial cell death in vitro and in murine experimental hydronephrosis.</a> Am J Pathol. 169, 388-399 (2006).</li><li>Henderson, N. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Galectin-3+expression+and+secretion+links+macrophages+to+the+promotion+of+renal+fibrosis.">Galectin-3 expression and secretion links macrophages to the promotion of renal fibrosis.</a> Am J Pathol. 172, 288-298 (2008).</li><li>Kitamoto, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+liposome+clodronate+on+renal+leukocyte+populations+and+renal+fibrosis+in+murine+obstructive+nephropathy.">Effects of liposome clodronate on renal leukocyte populations and renal fibrosis in murine obstructive nephropathy.</a> J Pharmacol Sci. 111, 285-292 (2009).</li><li>Tapmeier, T. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reimplantation+of+the+ureter+after+unilateral+ureteral+obstruction+provides+a+model+that+allows+functional+evaluation.">Reimplantation of the ureter after unilateral ureteral obstruction provides a model that allows functional evaluation.</a> Kidney Int. 73, 885-889 (2008).</li><li>Cochrane, A. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Renal+structural+and+functional+repair+in+a+mouse+model+of+reversal+of+ureteral+obstruction.">Renal structural and functional repair in a mouse model of reversal of ureteral obstruction.</a> J Am Soc Nephrol. 16, 3623-3630 (2005).</li><li>Puri, T. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+kidney+disease+induced+in+mice+by+reversible+unilateral+ureteral+obstruction+is+dependent+on+genetic+background.">Chronic kidney disease induced in mice by reversible unilateral ureteral obstruction is dependent on genetic background.</a> Am J Physiol Renal Physiol. 298, F1024-F1032 (2010).</li><li>Heikkila, J., Holmberg, C., Kyllonen, L., Rintala, R., Taskinen, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+risk+of+end+stage+renal+disease+in+patients+with+posterior+urethral+valves.">Long-term risk of end stage renal disease in patients with posterior urethral valves.</a> J Urol. 186, 2392-2396 (2011).</li><li>Ravanan, R., Tomson, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Natural+history+of+postobstructive+nephropathy:+a+single-center+retrospective+study.">Natural history of postobstructive nephropathy: a single-center retrospective study.</a> Nephron Clin Pract. 105, c165-c170 (2007).</li><li>Bascands, J. L., Schanstra, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Obstructive+nephropathy:+insights+from+genetically+engineered+animals.">Obstructive nephropathy: insights from genetically engineered animals.</a> Kidney Int. 68, 925-937 (2005).</li><li>Park, H. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Postobstructive+regeneration+of+kidney+is+derailed+when+surge+in+renal+stem+cells+during+course+of+unilateral+ureteral+obstruction+is+halted.">Postobstructive regeneration of kidney is derailed when surge in renal stem cells during course of unilateral ureteral obstruction is halted.</a> Am J Physiol Renal Physiol. 298, F357-F364 (2010).</li><li>Manson, S. R., Niederhoff, R. A., Hruska, K. A., Austin, P. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endogenous+BMP-7+is+a+critical+molecular+determinant+of+the+reversibility+of+obstruction-induced+renal+injuries.">Endogenous BMP-7 is a critical molecular determinant of the reversibility of obstruction-induced renal injuries.</a> Am J Physiol Renal Physiol. 301, F1293-F1302 (2011).</li><li>Manson, S. R., Niederhoff, R. A., Hruska, K. A., Austin, P. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+BMP-7-Smad1/5/8+pathway+promotes+kidney+repair+after+obstruction+induced+renal+injury.">The BMP-7-Smad1/5/8 pathway promotes kidney repair after obstruction induced renal injury.</a> J Urol. 185, 2523-2530 (2011).</li><li>Chaves, L. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contrasting+effects+of+systemic+monocyte/macrophage+and+CD4++T+cell+depletion+in+a+reversible+ureteral+obstruction+mouse+model+of+chronic+kidney+disease.">Contrasting effects of systemic monocyte/macrophage and CD4+ T cell depletion in a reversible ureteral obstruction mouse model of chronic kidney disease.</a> Clin Dev Immunol. 2013, 836-989 (2013).</li><li>Haque, M. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Longitudinal+changes+in+MRI+markers+in+a+reversible+unilateral+ureteral+obstruction+mouse+model:+preliminary+experience.">Longitudinal changes in MRI markers in a reversible unilateral ureteral obstruction mouse model: preliminary experience.</a> J Magn Reson Imaging. 39, 835-841 (2014).</li><li>Bai, Z. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Arterially+transplanted+mesenchymal+stem+cells+in+a+mouse+reversible+unilateral+ureteral+obstruction+model:+in+vivo+bioluminescence+imaging+and+effects+on+renal+fibrosis.">Arterially transplanted mesenchymal stem cells in a mouse reversible unilateral ureteral obstruction model: in vivo bioluminescence imaging and effects on renal fibrosis.</a> Chinese Med J. 126, 1890-1894 (2013).</li><li>Chaabane, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Renal+functional+decline+and+glomerulotubular+injury+are+arrested+but+not+restored+by+release+of+unilateral+ureteral+obstruction+(UUO).">Renal functional decline and glomerulotubular injury are arrested but not restored by release of unilateral ureteral obstruction (UUO).</a> Am J Physiol Renal Physiol. 304, F432-F439 (2013).</li><li>Thornhill, B. A., Burt, L. E., Chen, C., Forbes, M. S., Chevalier, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Variable+chronic+partial+ureteral+obstruction+in+the+neonatal+rat:+a+new+model+of+ureteropelvic+junction+obstruction.">Variable chronic partial ureteral obstruction in the neonatal rat: a new model of ureteropelvic junction obstruction.</a> Kidney Int. 67, 42-52 (2005).</li><li>Thornhill, B. A., Chevalier, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Variable+partial+unilateral+ureteral+obstruction+and+its+release+in+the+neonatal+and+adult+mouse.">Variable partial unilateral ureteral obstruction and its release in the neonatal and adult mouse.</a> Methods Mol Biol. 886, 381-392 (2012).</li></ol>

The authors have no competing or conflicting interests to disclose.

Obstruction of the kidney may affect native or transplanted kidneys and results in kidney injury and scarring. The R-UUO model is of relevance to human disease as patients often present with obstruction of the renal tract secondary to prostatic hypertrophy, posterior urethral valves etc. that requires decompression with patients commonly left with significant residual kidney impairment that has no current treatment options and may eventually lead to end stage kidney disease.8,9 The models detailed here enable ...

The overall aim of the experimental model described here is to induce obstructive nephropathy by unilateral ureteric obstruction (UUO), which can either be irreversible (UUO) or reversible (R-UUO). A simple irreversible model of UUO is presented in which the left ureter is permanently obstructed by ligation with a suture or by the application of a ligating clip. This results in marked dilatation of the ureter together with reduced renal blood flow and glomerular filtration. Renal histology demonstrates tubular dilatation and increasingly severe interstitial renal inflammation and fibrosis. Irreversible UUO is a useful model and has been adopted by many researchers in the study of both renal inflammation and fibrosis.1-4 Although the irreversible model of UUO requires some surgical expertise it is relatively straightforward and is often used to seek insights into the pathogenesis of interstitial renal injury and the ensuing fibrosis. Also presented is a less frequently used R-UUO model using a modification of the method originally described by Tapmeier et al.5 The R-UUO model has much future potential for the study of inflammatory and immune processes, cellular and tissue regeneration as well as the subsequent tissue remodeling following the removal of an injurious stimulus. 
The more challenging R-UUO model has been used by a limited number of investigators with some groups employing a significantly different surgical technique to that described here6,7 though with interesting results. In the R-UUO model presented, the ureter is ligated to induce complete ureteric obstruction for a period of time sufficient to induce the level of injury and fibrosis desired: 7 days of UUO was chosen in the method described here. The ureteric obstruction is reversed and the kidney is allowed to decompress for a period of time determined by the investigator before the mice are culled and the kidney removed for analysis: 7 days of decompression was chosen in the method described here though a longer period would be chosen if the resolution of inflammation and fibrosis was being studied. Although the method described here requires significant surgical expertise, it offers several advantages over other R-UUO models. The application of soft walled plastic tubing to the obstructed ureter prevents excessive ureteric dilation and this facilitates the subsequent manipulation and anastomosis of the ureter. Furthermore, in the described R-UUO model the ureter is divided thereby allowing the removal of any residual urinary sediment and debris. This confirms that the remaining ureter lumen is de-obstructed and patent prior to anastomosis to the bladder. 
Experiments incorporating both the irreversible and reversible UUO models can provide researchers with a powerful insight into the molecular and cellular mechanisms of both injury and subsequent resolution and regeneration. Thus, the model of R-UUO described here would be highly relevant to researchers interested in post-inflammatory tissue remodeling and how this can be modulated.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Blunt Dissecting Scissors</td>
			<td>Fine Science Tools</td>
			<td>14072-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Spring Scissors &#8211; straight</td>
			<td>Fine Science Tools</td>
			<td>15000-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Toothed forceps</td>
			<td>Fine Science Tools</td>
			<td>11021-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Angled forceps x 2</td>
			<td>Fine Science Tools</td>
			<td>00649-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight forceps</td>
			<td>Fine Science Tools</td>
			<td>00632-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Colibri 3cm wire retractor</td>
			<td>Fine Science Tools</td>
			<td>17000-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Castroviejo needle holder with lock</td>
			<td>Fine Science Tools</td>
			<td>12565-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Wound clip applicator</td>
			<td>Fine Science Tools</td>
			<td>12031-07</td>
			<td></td>
		</tr>
		<tr>
			<td>7mm wound clips</td>
			<td>Fine Science Tools</td>
			<td>12032-07</td>
			<td></td>
		</tr>
		<tr>
			<td>Castroviejo micro needle holder with lock</td>
			<td>Fine Science Tools</td>
			<td>12060-01</td>
			<td></td>
		</tr>
		<tr>
			<td>OPMI pico microscope</td>
			<td>Carl Zeiss</td>
			<td>S100</td>
			<td></td>
		</tr>
		<tr>
			<td>Heat electronic pad</td>
			<td>Cozee Comfort</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>6/O silk braided suture</td>
			<td>Harvard Apparatus</td>
			<td>72-3287</td>
			<td></td>
		</tr>
		<tr>
			<td>9/O Dafilon (polyamide) suture</td>
			<td>B-Braun</td>
			<td>G1111434</td>
			<td></td>
		</tr>
		<tr>
			<td>5/O braided silk suture</td>
			<td>Harvard Apparatus</td>
			<td>51-7680</td>
			<td></td>
		</tr>
		<tr>
			<td>Regular bevel needle, 1 inch, 21G</td>
			<td>Becton, Dickinson and Company</td>
			<td>305175</td>
			<td></td>
		</tr>
		<tr>
			<td>1 ml syringe slip tip</td>
			<td>Becton, Dickinson and Company</td>
			<td>300184</td>
			<td></td>
		</tr>
		<tr>
			<td>Wypall paper swabs</td>
			<td>Kimberley-Clark</td>
			<td>L40</td>
			<td>Sterilised (Autoclave)</td>
		</tr>
		<tr>
			<td>Cotton wool buds</td>
			<td>Johnson and Johnson</td>
			<td>n/a</td>
			<td>Sterilised (Autoclave)</td>
		</tr>
		<tr>
			<td>Plain drapes</td>
			<td>Guardian</td>
			<td>CB03</td>
			<td>Sterilised (Autoclave)</td>
		</tr>
		<tr>
			<td>Soft wall silicone rubber tubing</td>
			<td></td>
			<td></td>
			<td>Silicone tubing - internal diameter 1.0 mm; external diameter 2.0 mm; wall thickness 0.6 mm</td>
		</tr>
		<tr>
			<td>(Lacri-Lube) White soft paraffin 57.3%, mineral oil 42.5% and lanolin alcohols 0.2%</td>
			<td>Allergan Ltd</td>
			<td>21956GB10X</td>
			<td></td>
		</tr>
		<tr>
			<td>(Videne) Povidone-iodine 10%</td>
			<td>Ecolab Ltd</td>
			<td>PL 04509/0041</td>
			<td></td>
		</tr>
		<tr>
			<td>(Vetalar V) Ketamine hydrochloride</td>
			<td>Pfizer Animal Health</td>
			<td>Vm 42058/4165</td>
			<td>100mg/ml solution</td>
		</tr>
		<tr>
			<td>(Domitor) Medetomidine hydrochloride</td>
			<td>Orion Pharma</td>
			<td>Vm 06043/4003</td>
			<td>1mg/ml</td>
		</tr>
		<tr>
			<td>(Vetergesic) Bupernorphine hydrochloride</td>
			<td>Alsto Animal Health</td>
			<td>Vm 00063/4002</td>
			<td>0.3mg/ml</td>
		</tr>
		<tr>
			<td>(Antisedan) Atipamezole hydrochoride</td>
			<td>Orion Pharma</td>
			<td>Vm 06043/4004</td>
			<td>5mg/ml</td>
		</tr>
		<tr>
			<td>(Adept) 4% Icodextrin</td>
			<td>Baxter</td>
			<td></td>
			<td>Adhesion reduction solution</td>
		</tr>
		<tr>
			<td>NaCl 0.9%</td>
			<td>Baxter</td>
			<td>FKE1323</td>
			<td></td>
		</tr>
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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 appearance of the kidney changes markedly following ureteric obstruction and it becomes paler and tense to palpation with time (Figure 1). There is increasing dilatation of the ureter proximal to the obstruction and the renal pelvis. The kidney becomes increasingly atrophic as the duration of obstructive nephropathy increases with prolonged obstruction resulting in a marked thinning of the renal cortex and medulla. Following the reversal of the ureteric obstruction, the colour of the kidney becomes d...

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A Murine Model of Irreversible and Reversible Unilateral Ureteric Obstruction

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 ...

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30.2K Views.  University of Edinburgh. 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.

Video: A Murine Model of Irreversible and Reversible Unilateral Ureteric Obstruction

A Contusive Model of Unilateral Cervical Spinal Cord Injury Using the Infinite Horizon Impactor

While the majority of human spinal cord injuries occur in the cervical spinal cord, the vast majority of laboratory research employs animal models of spinal cord injury (SCI) in which the thoracic spinal cord is injured. Additionally, because most human cord injuries occur as the result of blunt, non-penetrating trauma (e.g. motor vehicle accident, sporting injury) where the spinal cord is violently struck by displaced bone or soft tissues, the majority of SCI researchers are of the opinion that the most clinically relevant injury models are those in which the spinal cord is rapidly contused.1 Therefore, an important step in the preclinical evaluation of novel treatments on their way to human translation is an assessment of their efficacy in a model of contusion SCI within the cervical spinal cord. Here, we describe the technical aspects and resultant anatomical and behavioral outcomes of an unilateral contusive model of cervical SCI that employs the Infinite Horizon spinal cord injury impactor.
Sprague Dawley rats underwent a left-sided unilateral laminectomy at C5. To optimize the reproducibility of the biomechanical, functional, and histological outcomes of the injury model, we contused the spinal cords using an impact force of 150 kdyn, an impact trajectory of 22.5&deg; (animals rotated at 22.5&deg;), and an impact location off of midline of 1.4 mm. Functional recovery was assessed using the cylinder rearing test, horizontal ladder test, grooming test and modified Montoya's staircase test for up to 6 weeks, after which the spinal cords were evaluated histologically for white and grey matter sparing.
The injury model presented here imparts consistent and reproducible biomechanical forces to the spinal cord, an important feature of any experimental SCI model. This results in discrete histological damage to the lateral half of the spinal cord which is largely contained to the ipsilateral side of injury. The injury is well tolerated by the animals, but does result in functional deficits of the forelimb that are significant and sustained in the weeks following injury. The cervical unilateral injury model presented here may be a resource to researchers who wish to evaluate potentially promising therapies prior to human translation.

A reliable and repeatable way to produce a cervical unilateral spinal cord injury using the Infinite Horizon impactor is described. The method takes advantage of a custom designed frame and clamp to stabilize the spine. The standardized procedure and biomechanical injury parameters result in sufficient and sustained injuries.

While the majority of human spinal cord injuries occur in the cervical spinal cord, the vast majority of laboratory research employs animal models of ...

A Murine Closed-chest Model of Myocardial Ischemia and Reperfusion

Surgical trauma by thoracotomy in open-chest models of coronary ligation induces an immune response which modifies different mechanisms involved in ischemia and reperfusion. Immune response includes cytokine expression and release or secretion of endogenous ligands of innate immune receptors. Activation of innate immunity can potentially modulate infarct size. We have modified an existing murine closed-chest model using hanging weights which could be useful for studying myocardial pre- and postconditioning and the role of innate immunity in myocardial ischemia and reperfusion. This model allows animals to recover from surgical trauma before onset of myocardial ischemia.
Volatile anesthetics have been intensely studied and their preconditioning effect for the ischemic heart is well known. However, this protective effect precludes its use in open chest models of coronary artery ligation. Thus, another advantage could be the use of the well controllable volatile anesthetics for instrumentation in a chronic closed-chest model, since their preconditioning effect lasts up to 72 hours. Chronic heart diseases with intermittent ischemia and multiple hit models are other possible applications of this model.
For the chronic closed-chest model, intubated and ventilated mice undergo a lateral blunt thoracotomy via the 4th intercostal space. Following identification of the left anterior descending a ligature is passed underneath the vessel and both suture ends are threaded through an occluder. Then, both suture ends are passed through the chest wall, knotted to form a loop and left in the subcutaneous tissue. After chest closure and recovery for 5 days, mice are anesthetized again, chest skin is reopened and hanging weights are hooked up to the loop under ECG control.
At the end of the ischemia/reperfusion protocol, hearts can be stained with TTC for infarct size assessment or undergo perfusion fixation to allow morphometric studies in addition to histology and immunohistochemistry.

Surgical trauma induces an inflammatory response. Cytokines and endogenous ligands are known to modulate myocardial infarct size following ischemia and reperfusion. We present a modified closed-chest model of murine ischemia and reperfusion using hanging weights to minimize effects of thoracotomy.

Surgical trauma by thoracotomy in open-chest models of coronary ligation induces an immune response which modifies different mechanisms involved in ...

A Murine Model of Muscle Training by Neuromuscular Electrical Stimulation

Neuromuscular electrical stimulation (NMES) is a common clinical modality that is widely used to restore1, maintain2 or enhance3-5 muscle functional capacity. Transcutaneous surface stimulation of skeletal muscle involves a current flow between a cathode and an anode, thereby inducing excitement of the motor unit and the surrounding muscle fibers. 
NMES is an attractive modality to evaluate skeletal muscle adaptive responses for several reasons. First, it provides a reproducible experimental model in which physiological adaptations, such as myofiber hypertophy and muscle strengthening6, angiogenesis7-9, growth factor secretion9-11, and muscle precursor cell activation12 are well documented. Such physiological responses may be carefully titrated using different parameters of stimulation (for Cochrane review, see 13). In addition, NMES recruits motor units non-selectively, and in a spatially fixed and temporally synchronous manner14, offering the advantage of exerting a treatment effect on all fibers, regardless of fiber type. Although there are specified contraindications to NMES in clinical populations, including peripheral venous disorders or malignancy, for example, NMES is safe and feasible, even for those who are ill and/or bedridden and for populations in which rigorous exercise may be challenging. 
Here, we demonstrate the protocol for adapting commercially available electrodes and performing a NMES protocol using a murine model. This animal model has the advantage of utilizing a clinically available device and providing instant feedback regarding positioning of the electrode to elicit the desired muscle contractile effect. For the purpose of this manuscript, we will describe the protocol for muscle stimulation of the anterior compartment muscles of a mouse hindlimb.

A murine model of neuromuscular electrical stimulation (NMES), a safe and inexpensive clinical modality, to the anterior compartment muscles is described. This model has the advantage of modifying a readily available clinical device for the purpose of eliciting targeted and specific muscle contractions in mice.

Neuromuscular electrical stimulation (NMES) is a common clinical modality that is widely used to restore1, maintain2 or enhance3-5 muscle functional ...

Murine Model of Wound Healing

Wound healing and repair are the most complex biological processes that occur in human life. After injury, multiple biological pathways become activated. Impaired wound healing, which occurs in diabetic patients for example, can lead to severe unfavorable outcomes such as amputation. There is, therefore, an increasing impetus to develop novel agents that promote wound repair. The testing of these has been limited to large animal models such as swine, which are often impractical. Mice represent the ideal preclinical model, as they are economical and amenable to genetic manipulation, which allows for mechanistic investigation. However, wound healing in a mouse is fundamentally different to that of humans as it primarily occurs via contraction. Our murine model overcomes this by incorporating a splint around the wound. By splinting the wound, the repair process is then dependent on epithelialization, cellular proliferation and angiogenesis, which closely mirror the biological processes of human wound healing. Whilst requiring consistency and care, this murine model does not involve complicated surgical techniques and allows for the robust testing of promising agents that may, for example, promote angiogenesis or inhibit inflammation. Furthermore, each mouse acts as its own control as two wounds are prepared, enabling the application of both the test compound and the vehicle control on the same animal. In conclusion, we demonstrate a practical, easy-to-learn, and robust model of wound healing, which is comparable to that of humans.

A murine model of cutaneous wound healing that can be used to assess therapeutic compounds in physiological and pathophysiological settings.

Wound healing and repair are the most complex biological processes that occur in human life. After injury, multiple biological pathways become activated. ...

A Murine Model of Subarachnoid Hemorrhage

In this video publication a standardized mouse model of subarachnoid hemorrhage (SAH) is presented. Bleeding is induced by endovascular Circle of Willis perforation (CWp) and proven by intracranial pressure (ICP) monitoring. Thereby a homogenous blood distribution in subarachnoid spaces surrounding the arterial circulation and cerebellar fissures is achieved. Animal physiology is maintained by intubation, mechanical ventilation, and continuous on-line monitoring of various physiological and cardiovascular parameters: body temperature, systemic blood pressure, heart rate, and hemoglobin saturation. Thereby the cerebral perfusion pressure can be tightly monitored resulting in a less variable volume of extravasated blood. This allows a better standardization of endovascular filament perforation in mice and makes the whole model highly reproducible. Thus it is readily available for pharmacological and pathophysiological studies in wild type and genetically altered mice.

A standardized mouse model of subarachnoid hemorrhage by intraluminal Circle of Willis perforation is described. Vessel perforation and subarachnoid bleeding are monitored by intracranial pressure monitoring. In addition various vital parameters are recorded and controlled to maintain physiologic conditions.

In this video publication a standardized mouse model of subarachnoid  hemorrhage (SAH) is presented. Bleeding is induced by endovascular Circle of  Willis ...

A Preclinical Murine Model of Hepatic Metastases

Numerous murine models have been developed to study human cancers and advance the understanding of cancer treatment and development. Here, a preclinical, murine pancreatic tumor model of hepatic metastases via a hemispleen injection of syngeneic murine pancreatic tumor cells is described. This model mimics many of the clinical conditions in patients with metastatic disease to the liver. Mice consistently develop metastases in the liver allowing for investigation of the metastatic process, experimental therapy testing, and tumor immunology research.

A preclinical, murine model of hepatic metastases performed via a hemispleen injection technique.

Numerous murine models have been developed to study human cancers and advance the understanding of cancer treatment and development. Here, a preclinical, ...

Pre-clinical Orthotopic Murine Model of Human Prostate Cancer

To study the multifaceted biology of prostate cancer, pre-clinical in vivo models offer a range of options to uncover critical biological information about this disease. The human orthotopic prostate cancer xenograft mouse model provides a useful alternative approach for understanding the specific interactions between genetically and molecularly altered tumor cells, their organ microenvironment, and for evaluation of efficacy of therapeutic regimens. This is a well characterized model designed to study the molecular events of primary tumor development and it recapitulates the early events in the metastatic cascade prior to embolism and entry of tumor cells into the circulation. Thus it allows elucidation of molecular mechanisms underlying the initial phase of metastatic disease. In addition, this model can annotate drug targets of clinical relevance and is a valuable tool to study prostate cancer progression. In this manuscript we describe a detailed procedure to establish a human orthotopic prostate cancer xenograft mouse model.

Prostate cancer is the second most common cause of cancer-related deaths in the United States. An orthotopic cancer model provides a useful approach to understand the biology of prostate cancer and to evaluate the efficacy of therapeutic regimens. This protocol describes detailed steps necessary to establish an orthotopic prostate cancer mouse model.

To study the multifaceted biology of prostate cancer, pre-clinical in vivo models offer a range of options to uncover critical biological information ...

Generating a Murine Orthotopic Metastatic Breast Cancer Model and Performing Murine Radical Mastectomy

In vivo mouse models to assess breast cancer progression are essential for cancer research, including preclinical drug developments. However, the majority of the practical and technical details are commonly omitted in published manuscripts which, therefore, makes it challenging to reproduce the models, particularly when it involves surgical techniques. Bioluminescence technology allows for the evaluation of small amounts of cancer cells even when a tumor is not palpable. Utilizing luciferase-expressing cancer cells, we establish a breast cancer orthotopic inoculation technique with a high tumorigenesis rate. Lung metastasis is assessed utilizing an ex vivo technique. We, then, establish a mastectomy model with a low local recurrence rate to assess the metastatic tumor burden. Herein, we describe, in detail, the surgical techniques of orthotopic implantation and mastectomy for breast cancer with a high tumorigenesis rate and low local recurrence rates, respectively, to improve breast cancer model efficiency.

We introduce a murine orthotopic breast cancer model and radical mastectomy model with bioluminescence technology to quantify the tumor burden to mimic human breast cancer progression.

In vivo mouse models to assess breast cancer progression are essential for cancer research, including preclinical drug developments. However, the majority ...

Isometric Contractility Measurement of the Mouse Mesenteric Artery Using Wire Myography

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and drugs. It is widely used in many studies on the physiological functions of vascular smooth muscle, the pathogenesis of vascular diseases such as hypertension, and the development of smooth muscle relaxant drugs. The mouse is a widely used model animal with a large pool of disease models and genetically modified strains. We introduced this method to measure the isometric contraction of mouse mesenteric artery in detail. A 1.4-mm segment of mouse mesenteric resistance artery was isolated and mounted on a myograph chamber by passing two steel wires through its lumen. After equilibration and normalization steps, the vessel segment was potentiated by a high-K+ solution twice prior to the contraction assay. As an example of the application of this method in drug development, we measured the relaxant effect of a novel natural substance, neoliensinine, isolated from a Chinese herb, embryos of the lotus seed (Nelumbo nucifera Gaertn.) on mouse mesenteric arteries. The vessel segments mounted on the myograph chamber were stimulated with a high-K+ solution. When the force tension reached a stable sustained phase, cumulative doses of neoliensinine were added to the chamber. We found that neoliensinine had a dose-dependent relaxant effect on smooth muscle contraction, thus suggesting that it bears potential activity against hypertension. In addition, as the vessel segment can survive at least 4 hours after mounting and maintain contractility induced by the high-K+ solution for many times, we suggest that the wire myograph system may be used for the time-consuming process of drug screening.

The wire myograph technique is used to investigate vascular smooth muscle functions and screen new drugs. We report a detailed protocol for measuring the isometric contractility of the mouse mesenteric artery and for screening new relaxants of vascular smooth muscle.

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and ...

A Reversible Silicon Oil-Induced Ocular Hypertension Model in Mice

Elevated intraocular pressure (IOP) is a well-documented risk factor for glaucoma. Here we describe a novel, effective method for consistently inducing stable IOP elevation in mice that mimics the post-operative complication of using silicone oil (SO) as a tamponade agent in human vitreoretinal surgery. In this protocol, SO is injected into the anterior chamber of the mouse eye to block the pupil and prevent inflow of aqueous humor. The posterior chamber accumulates aqueous humor and this in turn increases the IOP of the posterior segment. A single SO injection produces reliable, sufficient, and stable IOP elevation, which induces significant glaucomatous neurodegeneration. This model is a true replicate of secondary glaucoma in the eye clinic. To further mimic the clinical setting, SO can be removed from the anterior chamber to reopen the drainage pathway and allow inflow of aqueous humor, which is drained through the trabecular meshwork (TM) at the angle of the anterior chamber. Because IOP quickly returns to normal, the model can be used to test the effect of lowering IOP on glaucomatous retinal ganglion cells. This method is straightforward, does not require special equipment or repeat procedures, closely simulates clinical situations, and may be applicable to diverse animal species. However, minor modifications may be required.

Here, we present a protocol to induce ocular hypertension and glaucomatous neurodegeneration in mouse eyes by intracameral injection of silicone oil and the procedure for silicone oil removal from the anterior chamber to return elevated intraocular pressure to normal.

Elevated intraocular pressure (IOP) is a well-documented risk factor for glaucoma. Here we describe a novel, effective method for consistently inducing ...