We would like to acknowledge Drs. Robert Gereau, Henry Lai, and Lara Crock for their helpful assistance in setting up this system. We would also like to acknowledge funding sources for this work (BJK &#8211; International Association for the Study of Pain Early Career Research Grant by the Scan&#124;Design Foundation by INGER &#38; JENS BRUUN and the Duquesne University Hunkele Dreaded Disease Fund).

The following protocol has been approved by the Institutional Animal Care and Use Committee at Duquesne University and is consistent with guidelines from the National Institutes of Health.
1. Surgical Implantation of Wire Electrodes and Urinary Bladder Catheter Using Sterile Techniques&#160;(total time 5-10 min)
<ol>
	<li>The following items should be prepared prior to starting surgery:
	<ol>
		<li>Actively monitor and adjust body temperature to 37.5&#177;0.5 &#176;C using temperature monitoring system with a battery-operated heating pad and animal body temperature probe (Materials List) Connect automatic system to a low-wattage heat lamp for additional control of body temperature.</li>
		<li>Equipment needed for isoflurane anesthesia includes compressed air (e.g. 100% O2), isoflurane vaporizer, isoflurane, isoflurane induction chamber (for initial knockdown), nose cone, and exhaust system.</li>
		<li>Assemble the proper surgical instruments, IV catheter, and electrodes (Materials List)</li>
	</ol>
	</li>
	<li>Induce anesthetic state in induction chamber at 2-3% isoflurane. Remove mouse from chamber when righting reflex is lost (0-1 min).</li>
	<li>Place mouse in nose cone with 2-2.5% isoflurane. Pinch toe to check for full anesthesia. There should be no response from the animal. Start timer.</li>
	<li>Place 1 drop of ophthalmic ointment on each eye of mouse.</li>
	<li>Turn mouse over so dorsal side is facing down.</li>
	<li>Insert catheter through urethra and into bladder.</li>
	<li>Dip catheter in surgical lube to lubricate catheter.
	<ol>
		<li>Gently hold urethral opening with pointed forceps perpendicular to animal&#8217;s body. Holding catheter with the 2nd set of forceps, gently insert catheter end into urethra at an angle perpendicular to animal&#8217;s body.</li>
		<li>When catheter has gone into urethra ~1-2 mm, gently adjust catheter so that it is now parallel to the body aiming toward the head. This movement is necessary to avoid the pubic bone.</li>
		<li>Gently push catheter toward the mouse&#8217;s body. The catheter should slide into the body cavity smoothly and without resistance. Do not force catheter. If catheter does not enter smoothly, return to the perpendicular catheter position and start again.</li>
		<li>With catheter fully inserted in the bladder, gently push down on the mouse&#8217;s abdomen to expel bladder contents. Remove all urine from catheter by pushing a small piece of paper towel into catheter opening. When paper towel is soaked with urine, replace. Remove urine continuously throughout the remaining protocol including during bladder distention experiments (see Protocol&#160;3 below.)</li>
	</ol>
	</li>
	<li>Insert two silver wires into left abdominal muscles and one silver wire into chest (as a ground.)
	<ol>
		<li>Use alcohol or iodine to disinfect area lateral to the right of catheter (animal&#8217;s left side of body.)</li>
		<li>Expose left abdominal muscle.
		<ol>
			<li>Hold skin near left abdominal muscle with forceps. Make a small 1-2 cm incision with medium surgical scissors.</li>
			<li>Insert tip of scissors into incision and gently expand incision to 2 cm.</li>
			<li>Use forceps to move overlying fascia to expose superior oblique abdominal muscle. Be careful not to cut/rip any of the major blood vessels that are located in this region.</li>
		</ol>
		</li>
		<li>Bend three silver wires in half. Use sharp 21 G&#160;needle to take a small bite of the muscle. Then, push needle through muscle. When inserting in muscle be careful not to puncture any internal organs or induce unnecessary damage to the muscle.</li>
		<li>Stick a free end of one silver wire into the opening of the 21 G&#160;needle. Gently pull needle back through the muscle to draw the silver wire through the muscle bite. When needle is free from muscle, gently pull silver wire through muscle until the loop in the wire is flush with the muscle bite.</li>
		<li>Repeat steps 1.7.3-1.7.4 for second muscle wire. This wire should be placed ~0.5 cm from the first muscle wire. It is important that the two wires do not touch after insertion.</li>
		<li>Insert the third silver wire (ground) in the chest inferior to the heart.
		<ol>
			<li>Use sharp 21 G&#160;needle to take a small bite of skin (it is not necessary to make an incision or shave the skin) and push needle through the skin.</li>
			<li>Insert silver wire as described above and pull through skin.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Place a small amount of 37.5 &#176;C sterile mineral oil over the exposed muscles to keep them moist and pull skin over as much exposed muscle as possible.</li>
	<li>Inject 0.5 ml&#160;saline subcutaneously to help maintain fluid levels during subsequent steps.</li>
	<li>Gently roll animal onto side so that body temperature can be easily maintained.</li>
</ol>
2. Partial Anesthesia Step-down Procedure&#160;(total time ~75 min)
The following anesthesia protocol has been approved by the Institutional Animal Care and Use Committee at Duquesne University and is consistent with guidelines from the National Institutes of Health.
<ol>
	<li>Immediately after Section I (above) is complete, lower isoflurane to 1.5%. Hold at this level for 15 min.</li>
	<li>Lower isoflurane such that the animal is partially anesthetized.
	<ol>
		<li>Lower isoflurane by 0.125% every 15 min until animal exhibits a flexion response to toe pinch but does not vocalize or otherwise move.</li>
		<li>Typically, a level of 0.8-1.0% isoflurane is optimal, however levels can vary due to differences in anesthesia setups. Vocalization and/or ambulation are extremely rare events when anesthesia is at the proper level.</li>
	</ol>
	</li>
</ol>
3. Electromyogram (EMG) Recording During Bladder Distension with Graded Pressures (15-75 mmHg)&#160;(total time &#8211; variable depending on experiment)
<ol>
	<li>Setup the following items for EMG recording and bladder distention:
	<ol>
		<li>Setup an amplifier, a digitizer, and recording software. In this setup, the digitizer has one input for the amplifier and two inputs for the pressure regulator (pressure and stimulus marker inputs, see below section 3.1.2). Exact hardware and software setup is customizable (see Materials List for specific instructions).</li>
		<li>Setup a system to deliver specific bursts of air pressure to animal. This system is referred to as the &#8220;timed pressure regulator&#8221; in the following protocol. It allows for automated pressure delivery including digitization of air pressure, control of trial length, inter-trial interval automation, control of trial number, and stimulus onset digital signal (see Anderson&#160;et al.14 for schematic of a timed pressure regulator).</li>
	</ol>
	</li>
	<li>Connect all lead wires from mouse (2 abdominal wires + 1 ground wire) to the amplifier and the digitizer and begin recording EMG signal in digitizer software. Note what time &#8220;Write&#8221; is selected in digitizer software.</li>
	<li>Remove paper towel from bladder catheter. Connect catheter to air tubing (from timed pressure regulator.)</li>
	<li>Deliver a single 20 sec trial at 60 mmHg distention pressure.
	<ol>
		<li>Set the flow regulator to 60 mmHg (check pressure using an analog sphygmomanometer.)</li>
		<li>Set the timed pressure regulator to deliver a 20 sec prepressure interval (this will be the 2V stimulus signal recorded in digitizer software) followed by a 20 sec pressure distention pulse.</li>
		<li>Start the trial (prepressure interval + pressure distention trial). When the actual pressure trial begins, double check the pressure using the 3 sphygmomanometers connected to the system.</li>
		<li>During the pressure distention trial, a strong EMG signal (&#62;0.5 V) should be observed during and possibly after the 20 sec distention. The animal may exhibit abdominal movement but the animal should not vocalize or otherwise move. Although rare, if abnormal movements or vocalizations are observed, increase the isoflurane level.</li>
	</ol>
	</li>
	<li>If the 1st 60 mmHg distention produces an appropriate EMG signal, repeat step 3.4 two more times with an intertrial interval of 1-2 min.</li>
	<li>Upon receiving strong signals from the three 60 mmHg trials, perform experimental distentions.
	<ol>
		<li>Many studies use graded distensions starting at low pressure (10-15 mmHg) and working up to noxious pressures (75-80 mmHg) in 10-15 mmHg increments. See representative results below for example traces at different pressures.</li>
		<li>Typical trials include a 20 sec prepressure interval, a 20 sec pressure distention trial, and a 1-2 min intertrial interval.</li>
		<li>Perform three distentions at each pressure and then average across the three trials (to obtain a single visceromotor response for each pressure). In an alternative method, perform five distensions at each pressure, discard the high and low trials, and then average the remaining three responses.</li>
	</ol>
	</li>
	<li>Once experimental distensions are complete, stop recording, save the experiment file, and sacrifice animal using approved methods.</li>
</ol>
4. Analysis of Raw EMG Traces
<ol>
	<li>Analysis of visceromotor responses (i.e. EMG recordings) are easily done using on-board digitizer software or through a third-party program (Materials List) The following instructions are general instructions that could be used to analyze this data using multiple programs or using manual analysis.</li>
	<li>To export EMG, pressure, and stimulus signal data from digitizer software do the following:
	<ol>
		<li>Open experimental file.</li>
		<li>Click &#8220;File&#8221; &#8220;Export as.&#8221;</li>
		<li>Change the file type to &#8220;spreadsheet text (.txt),&#8221; name the file and click &#8220;Save.&#8221;</li>
		<li>In the pop-up window, change the output sample rate to &#8220;1000,&#8221; change the start time to the moment data collection began and change the end time to &#8220;MaxTime().&#8221; Select the box marking &#8220;time shift output so first line is a 0.0 sec,&#8221; and click &#8220;OK.&#8221;</li>
		<li>The data will now be exported as a text file that can be opened in a variety of programs. The file should contain four columns (Timer, VMR (EMG), Pressure, Stimulus).</li>
	</ol>
	</li>
	<li>For each experimental animal tested, perform the following mathematical operations.
	<ol>
		<li>Rectify EMG signal for the entire data set by calculating the absolute value of each time point.</li>
		<li>Determine the average EMG response during the background section of the experiment.</li>
		<li>Subtract this average background EMG response from every rectified data point in the experiment to obtain the background corrected data set.</li>
		<li>Determine the area under the curve (AUC) for each 20 sec prepressure interval in the background corrected data set.</li>
		<li>Determine the AUC for each 20 sec pressure distension trial in the background corrected data set.</li>
		<li>Divide each pressure distension trial AUC (obtained in step 4.3.5) by the lowest prepressure interval AUC from the whole experiment (obtained in step 4.3.4.). This is the normalized and background corrected AUC for each complete pressure distention trial.</li>
	</ol>
	</li>
	<li>For each experimental animal, average multiple normalized trials from a single pressure to obtain the normalized VMR in Volts*sec (Vsec).</li>
	<li>Perform the appropriate statistics on the data depending on the experimental design.</li>
</ol>

<ol>
	<li>Merskey, H., Bogduk, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Classification of Chronic Pain. , (1994).</li><li>Gureje, O., Korff, M. V., Simon, G. E., Gater, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Peristent+pain+and+well-being.+A+World+Health+Organization+study+in+primary+care.">Peristent pain and well-being. A World Health Organization study in primary care.</a> J. Am. Med. Assoc. 280, 147-151 (1998).</li><li>Gaskin, D. J., Richard, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+economic+costs+of+pain+in+the+United+States.">The economic costs of pain in the United States.</a> J. Pain. 13, 715-754 (2012).</li><li>Berry, S. H., 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=Prevalence+of+Symptoms+of+Bladder+Pain+Syndrome/Interstitial+Cystitis+Among+Adult+Females+in+the+United+States.">Prevalence of Symptoms of Bladder Pain Syndrome/Interstitial Cystitis Among Adult Females in the United States.</a> J. Urol. 186, 540-544 (2011).</li><li>Tunitsky, 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=Bladder+Pain+Syndrome/Interstitial+Cystitis+in+twin+sisters.">Bladder Pain Syndrome/Interstitial Cystitis in twin sisters.</a> J. Urol. 187, 148-152 (2012).</li><li>Tettamanti, G., 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=Influence+of+smoking,+coffee,+and+tea+consumption+on+Bladder+Pain+Syndrome+in+female+twins.">Influence of smoking, coffee, and tea consumption on Bladder Pain Syndrome in female twins.</a> Urology. 77, 1313-1317 (2011).</li><li>Warren, J. W., et al., Longo, D. 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=.">.</a> Harrison's Principles of Internal Medicine. , (2012).</li><li>Baldoni, F., Ercolani, M., Baldaro, B., Trombini, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stressful+events+and+psychological+symptoms+in+patients+with+functional+urinary+disorders.">Stressful events and psychological symptoms in patients with functional urinary disorders.</a> Perceptual Motor Skills. 80, 605-606 (1995).</li><li>Tannenbaum, C., Gray, M., Hoffstetter, S., Cardozo, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comorbidities+associated+with+bladder+dysfunction.">Comorbidities associated with bladder dysfunction.</a> Int. J. Clin. Practice. 67, 105-113 (2013).</li><li>Ness, T. J., Elhefni, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reliable+visceromotor+responses+are+evoked+by+noxious+bladder+distention+in+mice.">Reliable visceromotor responses are evoked by noxious bladder distention in mice.</a> J. Urol. 171, 1704-1708 (2004).</li><li>Ness, T. J., Richter, H. E., Varner, R. E., Fillingim, R. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+psychophysical+study+of+discomfort+produced+by+repeated+filling+of+the+urinary+bladder.">A psychophysical study of discomfort produced by repeated filling of the urinary bladder.</a> Pain. 76, 61-69 (1998).</li><li>Ness, T. J., Gebhart, G. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visceral+pain:+a+review+of+experimental+studies.">Visceral pain: a review of experimental studies.</a> Pain. 41, 167-234 (1990).</li><li>Crock, L. 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=Central+amygdala+metabotropic+glutamate+receptor+5+in+the+modulation+of+visceral+pain.">Central amygdala metabotropic glutamate receptor 5 in the modulation of visceral pain.</a> J. Neurosci. 32, 14217-14226 (2012).</li><li>Anderson, R. H., Ness, T. J., Gebhart, G. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+distension+control+device+useful+for+quantitative+studies+of+hollow+organ+sensation.">A distension control device useful for quantitative studies of hollow organ sensation.</a> Physiol. Behav. 41, 635-638 (1987).</li><li>Dimitrakov, J., 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=Pharmacologic+Management+of+Painful+Bladder+Syndrome/Interstitial+Cystitis:+A+Systematic+Review.">Pharmacologic Management of Painful Bladder Syndrome/Interstitial Cystitis: A Systematic Review.</a> Arch. Intern. Med. 167, 1922-1929 (2007).</li><li>Qin, C., Van Meerveld, B. G. r. e. e. n. w. o. o. d. -., Foreman, R. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spinal+neuronal+responses+to+urinary+bladder+stimulation+in+rats+with+corticosterone+or+aldosterone+onto+the+amygdala.">Spinal neuronal responses to urinary bladder stimulation in rats with corticosterone or aldosterone onto the amygdala.</a> J. Neurophysiol. 90, 2180-2189 (2003).</li><li>Randich, A., Mebane, H., DeBerry, J. J., Ness, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rostral+ventral+medulla+modulation+of+the+visceromotor+reflex+evoked+by+urinary+bladder+distension+in+female+rats.">Rostral ventral medulla modulation of the visceromotor reflex evoked by urinary bladder distension in female rats.</a> J. Pain. 9, 920-926 (2008).</li></ol>

The authors have nothing to disclose.

In humans, interstitial cystitis/bladder pain syndrome (IC/BPS) represents a significant clinical problem because patients have debilitating pain that is often unresponsive to regular pain treatment15. One of the major challenges in understanding and ultimately treating chronic bladder pain is to understand the normal neural processes that are involved in the response to noxious bladder distention. In order to overcome this challenge, an animal model of bladder pain is necessary. This model should be reproduci...

Chronic pain is officially defined as pain that persists for approximately three months, or for longer than the normal tissue healing time1. This type of pain is one of the main reasons that people are driven to seek medical attention2 and can cost up to $635 billion dollars annually3. Current chronic pain coping strategies are considered archaic; after decades of medical advancements, NSAIDs (nonsteroidal anti-inflammatory drugs) and opioids are still the primary treatments prescribed by physicians. However these treatments target all different types of pain in the same way, providing analgesic effects all over the body as opposed to specifically focusing on the cause of that exact pain incident. In order to better help those suffering from chronic pain, research attention should be shifted towards the etiology and possible pain-specific treatments associated with the most common causes of chronic pain. The goal of this manuscript is to describe a model used to better understand a condition known as Interstitial Cystitis/Painful Bladder Syndrome (IC/BPS).
IC/BPS is a debilitating condition that affects millions of people, primarily women over the age of 40 4. The exact causes of IC/BPS are not known, however prevalence has been linked to genetics5, specific diets, and high stress levels6. Symptoms of IC/BPS include but are not limited to: Increased urgency to urinate, increased frequency of urination, aching, piercing, or burning pain during bladder filling and voiding, and nocturia7. Patients experiencing these conditions have higher stress levels and are more anxious8. This increased stress results in increased pain, which augments the original stress levels. Studies have shown that depression and anxiety levels decrease following treatments that relieve urinary pain, thus effectively breaking this positive-feedback cycle9. One of the challenges in discovering new treatments for IC/BPS has been the paucity of a basic science understanding of bladder pain.
In order to study nervous system contributions to the condition, many animal models have been developed to mimic the pain and symptoms associated with IC/BPS. Traditionally, cystitis has been recreated in animals with the introduction of various chemicals such as mustard oil, acetone, lipopolysaccharide, hydrochloric acid, and cyclophosphamide into the bladder. However no foreign agents are present in the sterile urine of IC/BPS patients, thus questioning the validity of these models. Another murine model of urological pain is urinary bladder distension (UBD)10. In this model, compressed air of a specific pressure is delivered to the bladder of an awake or lightly anesthetized animal over a set period of time. Throughout the procedure, wires in the superior oblique abdominal muscles record electrical activity from the muscle (with an electromyogram (EMG)). This activity is known as the visceromotor response (VMR) and is reliable, reproducible measure of nociception10. Similar hollow organ distention (e.g. bladder, rectum) in healthy human volunteers produces feelings of discomfort and significant increases in reported pain11,12, traits that are often used to diagnose IC/BPS. Thus, in conjunction with electrical, pharmacological, and optogenetic methods of stimulation or inhibition, UBD is a useful and valid model for understanding both the peripheral and central nervous system components of bladder nociception and pain.
Here, we describe the procedure for UBD as is currently used in our laboratory within female members of the common mouse strain, C57Bl/6J. Due to difficulties in the catheterization process of males, this procedure is primarily performed in female mice. This protocol has been adapted from that developed by Ness et al.10 and previously published from our lab13. The following protocol describes four primary components of this model in anesthetized animals: (1) bladder catheter and recording electrode surgery (2) partial anesthesia induction (3) bladder distention and VMR recording, and (4) data analysis of raw VMR/EMG traces. Subtle differences in the UBD-VMR procedure due to choice of organism, anesthesia depth and induction, and body temperature are discussed below.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Infrared heating blanket and monitoring system</td>
			<td>Kent Scientific</td>
			<td>Right-Temp System&#160;</td>
			<td>This system is setup to monitor two separate temperatures.&#160; This should include the animal and the heating blanket.&#160; In addition, the system can automatically adjust the temperature to maintain a set temp.&#160; However, this automatic function produces electrical interference during EMG recording and must be turned off.&#160; Kent can provide a battery pack for the heating pad for use during the riEMG recording part of the experiment.</td>
		</tr>
		<tr>
			<td>Isoflurane vaporizer</td>
			<td>Draeger</td>
			<td>Vapor 19.1</td>
			<td>Any isoflurane vaporizer will work, but it is helpful to have one that has mutliple notches between 0 and 2% isoflurane.</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Multiple</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>100% Oxygen air tank</td>
			<td>Multiple</td>
			<td>n/a</td>
			<td>For ventilation of animal</td>
		</tr>
		<tr>
			<td>Air breathing grade</td>
			<td>Multiple</td>
			<td>n/a</td>
			<td>For bladder distention</td>
		</tr>
		<tr>
			<td>24GA 0.56inch IV catheter</td>
			<td>BD biosciences&#160;</td>
			<td>381411</td>
			<td>For bladder catheterization</td>
		</tr>
		<tr>
			<td>Surgilube (sterile)</td>
			<td>Savage Laboratories</td>
			<td>0281-0205-02</td>
			<td>Any surgical grade lubricate would work fine.</td>
		</tr>
		<tr>
			<td>Mineral oil (sterile)</td>
			<td>Multiple</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Saline (sterile)</td>
			<td>Multiple</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>AG8W Silver Wire, 2 m, 0.20 mm (.008&#34;) D, L, No Insulation&#160;</td>
			<td>Warner Instruments</td>
			<td>W4 64-1318&#160;</td>
			<td>Any silver wire with these specifications will work.&#160; Wire does not need to be &#34;chlorinated.&#34;</td>
		</tr>
		<tr>
			<td>opthalamic ointment</td>
			<td>Multiple</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Small surgical scissors</td>
			<td>Multiple</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Sharp forceps</td>
			<td>Multiple</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>21 Ga needle</td>
			<td>Multiple</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Grass amplifier P511 with 3-lead input cable</td>
			<td>Grass Instruments</td>
			<td>P511 (F-P5IC3/REV1)</td>
			<td>This is the &#34;amplifier&#34; used in the protocol.&#160; Amplifier with the following settings: Calibrator = 1 mV; Lo freq = 300 Hz; Amplification = 20; Hi freq = 10 kHz; Line filter = in.</td>
		</tr>
		<tr>
			<td>Cambridge Electronic Design (CED) 1401 Plus (or equivalent)</td>
			<td>Cambridge Electronic Design</td>
			<td>1401 Plus</td>
			<td>This is the &#34;digitizer&#34; used in the protocol.&#160; Other digitizer systems from WINDAQ or other companies would work fine; Need inputs for pressure signal, EMG, and stimulus signal.</td>
		</tr>
		<tr>
			<td>CED Spike2 software</td>
			<td>Cambridge Electronic Design</td>
			<td>Spike 2</td>
			<td>This is the the &#34;digitizer software&#34; used in the protocol.&#160; Should be from same manufacturer as digitizer.&#160; Program should be setup with 3 channels for pressure (0-100 mmHg scale), EMG signal (typically -5 to +5 V range), and stimulus marker (0-2 V) range.</td>
		</tr>
		<tr>
			<td>Flow chart from air tank to bladder catheter</td>
			<td>n/a</td>
			<td>n/a</td>
			<td>Sequence of connections from pressurized air tank to animal bladder:&#160; Air tank to 1/4 inch tubing to Gilmont flowmeter to Y connector.&#160; Branch 1 of y connector to to sphygmomanometer.&#160; Branch 2 to a single input on the 4-way gang valve to 4-way valve output to the timed pressure regulator to 3/32 tubing from timed pressure regulator to 2nd Y connector (branch 1 to sphygmomanometer) with branch 2 to 3/32 tubing&#160; to a 3rd Y connector.&#160; Branch 1 of Y connector to a 3rd sphygmomanometer and branch 2 to animal bladder catheter.</td>
		</tr>
		<tr>
			<td>Gilmont Flowmeter&#160;</td>
			<td>Gilmont</td>
			<td>GF8321-1401</td>
			<td>Multiple brands of flowmeters will work.&#160; For bladder distention air, flow meter should be able to handle high pressures (such as this Gilmont meter).&#160; For breathing air, flow rate should be adjustable down to 100 cc/min (typical mouse rate is 500-1000 cc/min).&#160;</td>
		</tr>
		<tr>
			<td>4-way gang valve</td>
			<td>Elite</td>
			<td></td>
			<td>This is a specific piece of equipment.&#160; The Elite gang valve is designed for fish tanks at low air pressures.&#160; In the bladder distention setup, this valve acts as a safety valves in case the pressure spikes.&#160; Too high pressure during initial turning on of the tank will ruin the pressure transducer in the Timed pressure regulator and/or the sphygnometers. In addition, by providing a small amount of leak in the system, this valve makes it easier to adjust the pressure between 10-80 mmHg.&#160;&#160;</td>
		</tr>
		<tr>
			<td>1/4 inch tubing</td>
			<td>Multiple</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>3/32 inch tubing</td>
			<td>Multiple</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>&#34;Y&#34; connectors (1/4 and 3/32 inch)</td>
			<td>Multiple</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Sphygmomanometer</td>
			<td>CVS</td>
			<td></td>
			<td>Any analog sphygmomanometer from a drug store will work for this application.&#160;&#160;</td>
		</tr>
		<tr>
			<td>Timed Pressure Regulator</td>
			<td>Custom</td>
			<td></td>
			<td>This is a custom built machine (Washington University in St. Louis Machine shop) that allows for automated pressure delivery including digitization of air pressure, control of trial length, inter-trial interval automation, control of trial number, and stimulus onset digital signal.&#160; However, the basic components of the system (pressure on and off for a given trial period) could be controlled with a simple on/off in-line switch.&#160;&#160; Such analog control of a trial would necessitate additional analysis parameters (see below section 4).&#160; In addition, one would have to manually assign the pressure based on the analog sphygmomanometer during data analysis.&#160;&#160;</td>
		</tr>
		<tr>
			<td>IGOR Pro</td>
			<td>Wavemetrics</td>
			<td>n/a</td>
			<td>For analysis of EMG signal. Many different types of software can be used for data analysis in these experiments.</td>
		</tr>
	</tbody>
</table>

urinary bladder distention evoked visceromotor responses as a model for bladder pain in mice

Approximately 3-8 million people in the United States suffer from interstitial cystitis/bladder pain syndrome (IC/BPS), a debilitating condition characterized by increased urgency and frequency of urination, as well as nocturia and general pelvic pain, especially upon bladder filling or voiding. Despite years of research, the cause of IC/BPS remains elusive and treatment strategies are unable to provide complete relief to patients. In order to study nervous system contributions to the condition, many animal models have been developed to mimic the pain and symptoms associated with IC/BPS. One such murine model is urinary bladder distension (UBD). In this model, compressed air of a specific pressure is delivered to the bladder of a lightly anesthetized animal over a set period of time. Throughout the procedure, wires in the superior oblique abdominal muscles record electrical activity from the muscle. This activity is known as the visceromotor response (VMR) and is a reliable and reproducible measure of nociception. Here, we describe the steps necessary to perform this technique in mice including surgical manipulations, physiological recording, and data analysis. With the use of this model, the coordination between primary sensory neurons, spinal cord secondary afferents, and higher central nervous system areas involved in bladder pain can be unraveled. This basic science knowledge can then be clinically translated to treat patients suffering from IC/BPS.

An illustration of the overall UBD-VMR setup can be seen in Figure 1A. Increasing pressure steps induce an increase in the raw VMR (i.e. EMG) (Figure 1B.) The raw VMR, pressure, and stimulus marker signal that should be observed during recording using digitizer software can be seen in Figure 2. The top channel seen in Figure 2 is the EMG trace (in V). The background signal of the EMG trace should be steady and low amplitude (&#60;0.2 V). If back...

Watch this Scientific Journal Video about Urinary Bladder Distention Evoked Visceromotor Responses as a Model for Bladder Pain in Mice at JoVE.com

Urinary Bladder Distention Evoked Visceromotor Responses as a Model for Bladder Pain in Mice

Approximately 3-8 million people in the United States suffer from interstitial cystitis/bladder pain syndrome (IC/BPS), a debilitating condition characterized in part by pelvic pain. In order to study nervous system contributions to the condition, a physiological model of urinary bladder pain is used in mice and rats.

Approximately 3-8 million people in the United States suffer from interstitial cystitis/bladder pain syndrome (IC/BPS), a debilitating condition ...

urinary-bladder-distention-evoked-visceromotor-responses-as-model-for

Research

JoVE Journal

Medicine

17.7K ビュー.  Duquesne University. 米国では約3から8000000人が間質性膀胱炎/膀胱痛症候群（IC / BPS）、骨盤痛によって部分的に特徴づけられる衰弱状態に苦しんでいます。条件神経系の寄与を検討するために、膀胱痛の生理学的モデルは、マウスおよびラットにおいて使用される。 

ビデオ: Urinary Bladder Distention Evoked Visceromotor Responses as a Model for Bladder Pain in Mice

Mouse Bladder Wall Injection

Mouse bladder wall injection is a useful technique to orthotopically study bladder phenomena, including stem cell, smooth muscle, and cancer biology. Before starting injections, the surgical area must be cleaned with soap and water and antiseptic solution. Surgical equipment must be sterilized before use and between each animal. Each mouse is placed under inhaled isoflurane anesthesia (2-5% for induction, 1-3% for maintenance) and its bladder exposed by making a midline abdominal incision with scissors. If the bladder is full, it is partially decompressed by gentle squeezing between two fingers. The cell suspension of interest is intramurally injected into the wall of the bladder dome using a 29 or 30 gauge needle and 1 cc or smaller syringe. The wound is then closed using wound clips and the mouse allowed to recover on a warming pad. Bladder wall injection is a delicate microsurgical technique that can be mastered with practice.

Mouse bladder wall injection is a useful approach to orthotopically study bladder stem cell and cancer biology. This delicate microsurgical method can be mastered with careful technique and practice.

Mouse bladder wall injection is a useful technique to orthotopically study bladder phenomena, including stem cell, smooth muscle, and cancer biology. ...

An Orthotopic Model of Murine Bladder Cancer

In this straightforward procedure, bladder tumors are established in female C57 mice through the use of catheterization, local cauterization, and subsequent cell adhesion. After their bladders are transurethrally catheterized and drained, animals are again catheterized to permit insertion of a platinum wire into bladders without damaging the urethra or bladder. The catheters are made of Teflon to serve as an insulator for the wire, which will conduct electrical current into the bladder to create a burn injury. An electrocautery unit is used to deliver 2.5W to the exposed end of the wire, burning away extracellular layers and providing attachment sites for carcinoma cells that are delivered in suspension to the bladder through a subsequent catheterization. Cells remain in the bladder for 90 minutes, after which the catheters are removed and the bladders allowed to drain naturally. The development of tumor is monitored via ultrasound. Specific attention is paid to the catheterization technique in the accompanying video.

Bladder tumors are established in female mice in a minimally invasive fashion through catheterization, local cauterization, and subsequent adhesion of carcinoma cells to the burn sites.

In this straightforward procedure, bladder tumors are established in female C57 mice through the use of catheterization, local cauterization, and ...

The Use of Cystometry in Small Rodents: A Study of Bladder Chemosensation

The lower urinary tract (LUT) functions as a dynamic reservoir that is able to store urine and to efficiently expel it at a convenient time. While storing urine, however, the bladder is exposed for prolonged periods to waste products. By acting as a tight barrier, the epithelial lining of the LUT, the urothelium, avoids re-absorption of harmful substances. Moreover, noxious chemicals stimulate the bladder's nociceptive innervation and initiate voiding contractions that expel the bladder's contents. Interestingly, the bladder's sensitivity to noxious chemicals has been used successfully in clinical practice, by intravesically infusing the TRPV1 agonist capsaicin to treat neurogenic bladder overactivity1. This underscores the advantage of viewing the bladder as a chemosensory organ and prompts for further clinical research. However, ethical issues severely limit the possibilities to perform, in human subjects, the invasive measurements that are necessary to unravel the molecular bases of LUT clinical pharmacology. A way to overcome this limitation is the use of several animal models2. Here we describe the implementation of cystometry in mice and rats, a technique that allows measuring the intravesical pressure in conditions of controlled bladder perfusion.
	After laparotomy, a catheter is implanted in the bladder dome and tunneled subcutaneously to the interscapular region. Then the bladder can be filled at a controlled rate, while the urethra is left free for micturition. During the repetitive cycles of filling and voiding, intravesical pressure can be measured via the implanted catheter. As such, the pressure changes can be quantified and analyzed. Moreover, simultaneous measurement of the voided volume allows distinguishing voiding contractions from non-voiding contractions3.
	Importantly, due to the differences in micturition control between rodents and humans, cystometric measurements in these animals have only limited translational value4. Nevertheless, they are quite instrumental in the study of bladder pathophysiology and pharmacology in experimental pre-clinical settings. Recent research using this technique has revealed the key role of novel molecular players in the mechano- and chemo-sensory properties of the bladder.

Cystometry is an efficient technique to measure bladder function of small animals in vivo. The bladder is continuously infused at rates controlled through an intravesical catheter, whereas the urethra is left free for micturition. This allows for repetitive filling and emptying of the bladder, while intravesical pressure and voided volume are recorded.

	The lower urinary tract (LUT) functions as a  dynamic reservoir that is able to store urine and to efficiently expel it at a  convenient time. While ...

An Orthotopic Bladder Tumor Model and the Evaluation of Intravesical saRNA Treatment

We present a novel method for treating bladder cancer with intravesically delivered small activating RNA (saRNA) in an orthotopic xenograft mouse bladder tumor model. The mouse model is established by urethral catheterization under inhaled general anesthetic. Chemical burn is then introduced to the bladder mucosa using intravesical silver nitrate solution to disrupt the bladder glycosaminoglycan layer and allows cells to attach. Following several washes with sterile water, human bladder cancer KU-7-luc2-GFP cells are instilled through the catheter into the bladder to dwell for 2 hours. Subsequent growth of bladder tumors is confirmed and monitored by in vivo bladder ultrasound and bioluminescent imaging. The tumors are then treated intravesically with saRNA formulated in lipid nanoparticles (LNPs). Tumor growth is monitored with ultrasound and bioluminescence. All steps of this procedure are demonstrated in the accompanying video.

Establishing an orthotopic bladder tumor model to evaluate antitumor effects of intravesically delivered saRNA and monitoring tumor growth by ultrasound and bioluminescent imaging.

We present a novel method for treating bladder cancer with intravesically delivered small activating RNA (saRNA) in an orthotopic xenograft mouse bladder ...

An Orthotopic Bladder Cancer Model for Gene Delivery Studies

Bladder cancer is the second most common cancer of the urogenital tract and novel therapeutic approaches that can reduce recurrence and progression are needed. The tumor microenvironment can significantly influence tumor development and therapy response. It is therefore often desirable to grow tumor cells in the organ from which they originated. This protocol describes an orthotopic model of bladder cancer, in which MB49 murine bladder carcinoma cells are instilled into the bladder via catheterization. Successful tumor cell implantation in this model requires disruption of the protective glycosaminoglycan layer, which can be accomplished by physical or chemical means. In our protocol the bladder is treated with trypsin prior to cell instillation. Catheterization of the bladder can also be used to deliver therapeutics once the tumors are established. This protocol describes the delivery of an adenoviral construct that expresses a luciferase reporter gene. While our protocol has been optimized for short-term studies and focuses on gene delivery, the methodology of mouse bladder catheterization has broad applications.

Implantation of cancer cells into the organ of origin can serve as a useful preclinical model to evaluate novel therapies. MB49 bladder carcinoma cells can be grown within the bladder following intravesical instillation. This protocol demonstrates catheterization of the mouse bladder for the purpose of tumor implantation and adenoviral delivery.

Bladder cancer is the second most common cancer of the urogenital tract and novel therapeutic approaches that can reduce recurrence and progression are ...

Induction of Invasive Transitional Cell Bladder Carcinoma in Immune Intact Human MUC1 Transgenic Mice:  A Model for Immunotherapy Development

A preclinical model of invasive bladder cancer was developed in human mucin 1 (MUC1) transgenic (MUC1.Tg) mice for the purpose of evaluating immunotherapy and/or cytotoxic chemotherapy. To induce bladder cancer, C57BL/6 mice (MUC1.Tg and wild type) were treated orally with the carcinogen N-butyl-N-(4-hydroxybutyl)nitrosamine (OH-BBN) at 3.0 mg/day, 5 days/week for 12 weeks. To assess the effects of OH-BBN on serum cytokine profile during tumor development, whole blood was collected via submandibular bleeds prior to treatment and every four weeks. In addition, a MUC1-targeted peptide vaccine and placebo were administered to groups of mice weekly for eight weeks. Multiplex fluorometric microbead immunoanalyses of serum cytokines during tumor development and following vaccination were performed. At termination, interferon gamma (IFN-&#947;)/interleukin-4 (IL-4) ELISpot analysis for MUC1 specific T-cell immune response and histopathological evaluations of tumor type and grade were performed. The results showed that: (1) the incidence of bladder cancer in both MUC1.Tg and wild type mice was 67%; (2) transitional cell carcinomas (TCC) developed at a 2:1 ratio compared to squamous cell carcinomas (SCC); (3) inflammatory cytokines increased with time during tumor development; and (4) administration of the peptide vaccine induces a Th1-polarized serum cytokine profile and a MUC1 specific T-cell response. All tumors in MUC1.Tg mice were positive for MUC1 expression, and half of all tumors in MUC1.Tg and wild type mice were invasive. In conclusion, using a team approach through the coordination of the efforts of pharmacologists, immunologists, pathologists and molecular biologists, we have developed an immune intact transgenic mouse model of bladder cancer that expresses hMUC1.

An N-butyl-N-(4-hydroxybutyl)nitrosamine-induced bladder cancer model was developed in human mucin 1 (MUC1) transgenic mice for the purpose of testing MUC1-directed immunotherapy. After administering a MUC1-targeted peptide vaccine, a cytotoxic T lymphocyte response to MUC1 was confirmed by measuring serum cytokine levels and T-cell specific activity.

A preclinical model of invasive bladder cancer was developed in human mucin 1 (MUC1) transgenic (MUC1.Tg) mice for the purpose of evaluating immunotherapy ...

Minimally Invasive Establishment of Murine Orthotopic Bladder Xenografts

Orthotopic bladder cancer xenografts are the gold standard to study molecular cellular manipulations and new therapeutic agents&#160;in vivo. Suitable cell lines are inoculated either by intravesical instillation (model of nonmuscle invasive growth) or intramural injection into the bladder wall (model of invasive growth). Both procedures are complex and highly time-consuming. Additionally, the superficial model has its shortcomings due to the lack of cell lines that are tumorigenic following instillation. Intramural injection, on the other hand, is marred by the invasiveness of the procedure and the associated morbidity for the host mouse.
With these shortcomings in mind, we modified previous methods to develop a minimally invasive approach for creating orthotopic bladder cancer xenografts. Using ultrasound guidance we have successfully performed percutaneous inoculation of the bladder cancer cell lines UM-UC1, UM-UC3 and UM-UC13 into 50 athymic nude. We have been able to demonstrate that this approach is time efficient, precise and safe. With this technique, initially a space is created under the bladder mucosa with PBS, and tumor cells are then injected into this space in a second step. Tumor growth is monitored at regular intervals with bioluminescence imaging and ultrasound. The average tumor volumes increased steadily in in all but one of our 50 mice over the study period.
In our institution, this novel approach, which allows bladder cancer xenograft inoculation in a minimally-invasive, rapid and highly precise way, has replaced the traditional model.

The established technique to inoculate primary invasive orthotopic bladder cancer xenografts requires laparotomy and mobilization of the bladder. This procedure inflicts significant morbidity on the mice, is technically challenging and time-consuming. We therefore developed a high-precision, percutaneous approach utilizing ultrasound guidance.

Orthotopic bladder cancer xenografts are the gold standard to study molecular cellular manipulations and new therapeutic agents&#160;in vivo. Suitable ...

The Sciatic Nerve Cuffing Model of Neuropathic Pain in Mice

Neuropathic pain arises as a consequence of a lesion or a disease affecting the somatosensory system. This syndrome results from maladaptive changes in injured sensory neurons and along the entire nociceptive pathway within the central nervous system. It is usually chronic and challenging to treat. In order to study neuropathic pain and its treatments, different models have been developed in rodents. These models derive from known etiologies, thus reproducing peripheral nerve injuries, central injuries, and metabolic-, infectious- or chemotherapy-related neuropathies. Murine models of peripheral nerve injury often target the sciatic nerve which is easy to access and allows nociceptive tests on the hind paw. These models rely on a compression and/or a section. Here, the detailed surgery procedure for the &#34;cuff model&#34; of neuropathic pain in mice is described. In this model, a cuff of PE-20 polyethylene tubing of standardized length (2 mm) is unilaterally implanted around the main branch of the sciatic nerve. It induces a long-lasting mechanical allodynia, i.e., a nociceptive response to a normally non-nociceptive stimulus that can be evaluated by using von Frey filaments. Besides the detailed surgery and testing procedures, the interest of this model for the study of neuropathic pain mechanism, for the study of neuropathic pain sensory and anxiodepressive aspects, and for the study of neuropathic pain treatments are also discussed.

Neuropathic pain is a consequence of a lesion or disease affecting the somatosensory system. The &#8220;cuff model&#8221; of neuropathic pain in mice consists of the implantation of a polyethylene cuff around the main branch of the sciatic nerve. Mechanical allodynia is tested using von Frey filaments.

Neuropathic pain arises as a consequence of a lesion or a disease affecting the somatosensory system. This syndrome results from maladaptive changes in ...

Bladder Smooth Muscle Strip Contractility as a Method to Evaluate Lower Urinary Tract Pharmacology

We describe an in vitro method to measure bladder smooth muscle contractility, and its use for investigating physiological and pharmacological properties of the smooth muscle as well as changes induced by pathology. This method provides critical information for understanding bladder function while overcoming major methodological difficulties encountered in in vivo experiments, such as surgical and pharmacological manipulations that affect stability and survival of the preparations, the use of human tissue, and/or the use of expensive chemicals. It also provides a way to investigate the properties of each bladder component (i.e. smooth muscle, mucosa, nerves) in healthy and pathological conditions.
The urinary bladder is removed from an anesthetized animal, placed in Krebs solution and cut into strips. Strips are placed into a chamber filled with warm Krebs solution. One end is attached to an isometric tension transducer to measure contraction force, the other end is attached to a fixed rod. Tissue is stimulated by directly adding compounds to the bath or by electric field stimulation electrodes that activate nerves, similar to triggering bladder contractions in vivo. We demonstrate the use of this method to evaluate spontaneous smooth muscle contractility during development and after an experimental spinal cord injury, the nature of neurotransmission (transmitters and receptors involved), factors involved in modulation of smooth muscle activity, the role of individual bladder components, and species and organ differences in response to pharmacological agents. Additionally, it could be used for investigating intracellular pathways involved in contraction and/or relaxation of the smooth muscle, drug structure-activity relationships and evaluation of transmitter release.
The in vitro smooth muscle contractility method has been used extensively for over 50 years, and has provided data that significantly contributed to our understanding of bladder function as well as to pharmaceutical development of compounds currently used clinically for bladder management.

This manuscript presents a simple, yet powerful, in vitro method for evaluating smooth muscle contractility in response to pharmacological agents or nerve stimulation. Main applications are drug screening and understanding tissue physiology, pharmacology,&#160;and pathology.

We describe an in vitro method to measure bladder smooth muscle contractility, and its use for investigating physiological and pharmacological properties ...

Evaluation of the Efficacy of the H. pylori Protein HP-NAP as a Therapeutic Tool for Treatment of Bladder Cancer in an Orthotopic Murine Model

Bladder cancer is one of the most common malignancies of the urogenital tract. Intravesical injection of Bacillus Calmette-Gu&#233;rin (BCG) is the gold standard treatment for the high-grade non-muscle invasive bladder cancer (NMIBC). However, since the treatment-related side effects are relevant, newer biological response modifiers with a better benefit/side effects ratio are needed.
The tumour microenvironment can influence both tumour development and therapy efficacy. In order to obtain a good model, it is desirable to implant tumour cells in the organ from which the cancer originates.
In this protocol, we describe a method for establishing a tumour in the bladder cavity of female mice and subsequent delivery of therapeutic agents; the latter are exemplified by our use of Helicobacter pylori neutrophil activating protein (HP-NAP). A preliminary chemical burn of the mucosa, followed by the injection of mouse urothelial carcinoma cell line MB49 via urethral catheterization, enables the cells to attach to the bladder mucosa. After a period, required to allow an initial proliferation of the cells, mice are treated with HP-NAP, administrated again via catheterization. The anti-tumour activity of HP-NAP is evaluated comparing the tumour volume, the extent of necrosis and the degree of vascularization between vehicle- and HP-NAP-treated animals.

Evaluation of the Efficacy of the H. pylori Protein HP-NAP as a Therapeutic Tool for Treatment of Bladder Cancer in an Orthotopic Murine Model

Here the method to establish a syngeneic mouse model of orthotopic bladder tumour to evaluate the anti-tumour efficacy of the bacterial protein HP-NAP is described.

Bladder cancer is one of the most common malignancies of the urogenital tract. Intravesical injection of Bacillus Calmette-Gu&#233;rin (BCG) is the gold ...