This study was funded by the Department of Surgery University of Vermont, Danish Council for Independent Research, and by the Odense University Hospital.

Animals were housed in the University of Vermont Animal Care Facility according to institutional guidelines. All animal experiments were carried out in accordance with the National Institutes of Health guide for the care and use of laboratory animals.
1. Intravesical Tube Implantation
<ol>
	<li>Preparation of tubing and instruments for the surgical procedure
	<ol>
		<li>Cut a 7-cm piece of PE10 tubing to make the catheter for implantation.</li>
		<li>Create a flare at one end of the PE10 tube by slowly advancing the end towards an open flame. 
		NOTE: Quickly withdraw the tube as soon as the flare develops.</li>
		<li>Apply three drops of all-purpose hot glue, using the low heat setting on a glue gun, at 4.5, 5, and 5.5 cm from the flared end on the outside of the PE10 tube. These will help secure the tube at the animal&#39;s back. (Figure 1)</li>
		<li>Sterilize the tubing by soaking it in 70% ethanol and then flush it with sterile 0.9% NaCl prior to use. Leave the tube filled to avoid introducing air bubbles into the system.</li>
		<li>Create a 30-gauge plug to seal the end of the PE10 catheter by separating a 30-gauge needle from the hub by manually manipulating the proximal end side-to-side. Apply a drop of hot glue to the end. Ensure that the seal is watertight. (Figure 2)</li>
		<li>Use the following microsurgical instruments: Two pairs of Dumont #7 curved microforceps, two pairs of Dumont #5 curved microforceps, a 21 G needle, ultrafine straight hemostat, micro scissors, small dissecting scissors, and a micro needle holder.</li>
		<li>Sterilize all instruments before starting the procedure.</li>
	</ol>
	</li>
	<li>Preparation of the animal
	<ol>
		<li>After anesthetizing the animal, shave the lower half of the abdomen first, then turn the animal prone and shave and clean the area on the upper back with 70% alcohol followed by betadine. Apply vet ointment to the eyes to prevent dryness. Next, use a pair of straight, blunt scissors and pair of Dumont #7 curved microforceps to make a 1.5 cm long skin incision between the scapulae and place the animal supine on top of a heating pad (37 &#176;C) covered with sterile drapes.</li>
		<li>Finally, clean the abdomen with alcohol and betadine.</li>
	</ol>
	</li>
	<li>Surgical procedure 
	NOTE: Perform all surgical procedures under an operating microscope with magnification ranging from 3.15X to 20X. After placing the animal onto the sterile drapes, put on sterile gloves. Continue using sterile procedures throughout the entire surgery.
	<ol>
		<li>Place the animal in an induction box and anesthetize using 2% inhaled isoflurane with an oxygen carrier (1 L/min). Maintain anesthesia throughout the procedure by placing the animal&#39;s head in a nose cone and using 2% inhaled isoflurane with an oxygen carrier (1 L/min). Begin the surgery after receiving a negative response from the toe-pinch test.</li>
		<li>Use a pair of straight, blunt scissors and pair of Dumont #7 curved microforceps to make a 1.5 cm lower, midline abdominal incision through the skin. Subsequently, create a matching incision through the fascia along the linea alba and muscle to expose the dome and upper half of the urinary bladder. Avoid injuring the bladder by applying upward traction to each tissue layer using a pair of Dumont #7 curved microforceps. Keep the abdominal viscera from desiccation by adding drops of warm physiological saline.</li>
		<li>Rotate the animal onto its side to access the incision on the nape of the neck. Push a narrow hemostat subcutaneously though the incision. The subcutaneous channel should start on the back, and continue along the side.</li>
		<li>Once the tip of the instrument reaches the bottom of the rib cage, turn the tip towards the midline and inside of the abdomen (there will be a slight pop when piercing the abdominal wall muscles). Continue advancing the hemostat until the tip is exposed at the abdominal incision under the muscular layer. (Figure 3)</li>
		<li>Grasp the &#34;non-flared&#34; end of the tubing with the hemostat and slowly retract the tool, pulling the end of the tube out via the incision at the back of the neck. Adjust the flared end of the tubing so that it lies directly above the dome of the bladder.</li>
		<li>Make a loose tie of 6-0 monofilament suture (non-absorbable) and place it on top of the bladder dome. This tie will be used later to secure the tube in the bladder.</li>
		<li>Place a small roll of lint-free tissue in the abdomen and behind the bladder to help stabilize and elevate it.</li>
		<li>Prepare to insert the flared end of the PE10 catheter into the bladder.
		<ol>
			<li>In the non-dominant hand, hold the dome of the bladder with Dumont #7 curved microforceps and maintain this grip until the catheter is placed in the bladder.</li>
			<li>Use a 21-gauge needle to make a cystotomy in the apex of the dome. Gently probe the cystotomy with a closed pair of #5 curved microforceps to make sure the catheter can easily pass through the hole.</li>
			<li>While still holding the bladder dome in the non-dominant hand, place the flared end of the PE10 catheter into the bladder (push the flare down to the bladder neck so that it does not slip out while securing it).</li>
			<li>Tie the 6-0 monofilament suture around the dome of the bladder and tubing with the tie placed anterior to the tubing. Be sure to tie the suture as high up on the bladder as possible to avoid artificially reducing bladder capacity. (Figure 4)</li>
			<li>Alternatively, secure the catheter using a purse string suture as follows. Make a loose purse string suture on the dome of the bladder using 6-0 monofilament. Follow steps 1.3.8.1 - 1.3.8.3 to perform the cystotomy and insert the catheter. Secure the tube by tying the purse string suture. (Figure 5)</li>
		</ol>
		</li>
		<li>Test the patency and seal of the tube in the bladder by attaching a 0.5 mL insulin syringe with a 30-gauge needle to the distal end of the tube. Slowly fill the bladder with 0.1 - 0.2 mL of 0.9% NaCl until a drop appears at the urethral orifice, then empty the bladder by aspiration. It is important that the bladder can be both filled and emptied.</li>
		<li>If no leaks occur at the dome, brace the bladder with a pair of curved microforceps and gently pull on the tubing until the flare is resting against the inside of the bladder dome.</li>
		<li>Before closing, remove the small roll of tissue, and make sure that the bladder is in its normal position.</li>
		<li>Close the abdominal wall in two layers (muscle and skin) with 6-0 running suture. It is preferable to approximate the rectus abdominis muscle by suturing only the edges of the anterior abdominal fascia (anterior wall of the rectus sheath).</li>
		<li>To secure the tubing at the animals back, gently rotate the animal onto its abdomen. Insert the subcutaneous portion of the metal anchor into the interscapular incision. (Figure 12) Use a 6-0 suture to secure the tube and anchor by encircling them with a vertical mattress suture.</li>
		<li>Make sure a glue bubble remains above and below the skin to prevent the tube from pulling out. Cut the tube approximately 2 cm above the skin.</li>
		<li>Gently insert the 30-gauge plug (step 1.1.5) into the end of the tube to prevent urine from leaking out.</li>
	</ol>
	</li>
	<li>Inject 0.5 mL 0.9% NaCl subcutaneously for hydration. Give post-operative analgesia immediately following surgery and maintain for 48 h.
	<ol>
		<li>Place the animal back into its cage located under an infrared lamp. Maintain constant observation until the animal moves around the cage freely.</li>
	</ol>
	</li>
	<li>Monitor the animal daily and allow it to recover for 5 days before recording.</li>
</ol>
2. Awake Cystometry Recording
<ol>
	<li>Preparation of the recording program, pressure transducer, and infusion pump.
	<ol>
		<li>Before anesthetizing the animal, connect the infusion pump, pressure transducer, and 22 G swivel using PE50 tubing. (Figure 6)</li>
		<li>Open the recording program (see table of materials for an example), on a computer to calibrate the system pressure and prepare for recording. Make sure to use the same settings during calibration and recording.
		<ol>
			<li>Fill a 20 mL syringe with 10 - 15 mL of room temperature 0.9% NaCl and load into the infusion pump. Program the pump to infuse at a rate of 0.6 mL/h.</li>
			<li>Secure the pressure transducer at the same height as the animal&#39;s bladder or the bottom of the recording cage.</li>
			<li>Attach the 22-gauge swivel to the end of the pressure transducer (PE50 - tubing - pressure transducer to swivel) 
			NOTE: The swivel is used to prevent the tube from twisting or kinking as the animal moves.</li>
			<li>Advance the syringe pump to flush 0.9% NaCl through the system. Make sure to remove all air bubbles before calibrating.</li>
			<li>With the recording program running, use a ruler to calibrate the pressure (cm/H2O). Slowly move the end of the PE50 tether from 0 to 30 cm. Adjust the zero if needed. 
			NOTE: The 0-cm mark should be at the same height as the floor of the recording cage and pressure transducer.</li>
		</ol>
		</li>
		<li>Suspend the 22-gauge swivel above the center of the recording cage. Ensure that the cage bottom allows urine to fall onto the collecting device of the balance positioned below the cage. Adjust the height of the tether so the mouse can freely move around the cage without straining or stretching the tubing. (Figure 7)</li>
		<li>When finished, check to make sure the system and external PE50 tubing are full of 0.9% NaCl and all air bubbles have been removed.</li>
	</ol>
	</li>
	<li>Preparation of the animal for recording
	<ol>
		<li>Anesthetize the animal with 2% inhaled isoflurane and place it on its abdomen. Remove the 30-gauge plug and slide the PE10 tubing (bladder catheter) into the end of the PE50 tether. Use hot glue to form a watertight seal.</li>
		<li>Turn off the anesthesia and place the animal in the recording cagewith a parallel-wire floor, which will allow urine to fall directly onto a collecting device placed on top of an analytical balance. (Figure 7)</li>
		<li>Start the recording once the animal is in the cage, but do not start infusing. Monitor the animal until it fully recovers from the anesthesia. Once the bladder pressure stabilizes, begin infusing 0.9% NaCl at a rate of 0.6 mL/h. 
		NOTE: Make a note in the recording program when any changes are made. It is important to have a record of when infusion starts, stops, or irregularities occur.</li>
		<li>Check the system for leaks and make sure the animal has easy access to food and water.</li>
		<li>Continue recording in a quiet room until three reproducible micturition cycles are obtained. 
		NOTE: The animal should be completely undisturbed throughout the recording. Preferably, use remote video monitoring to observe behavior.</li>
	</ol>
	</li>
</ol>

<ol><li>Perez, L. M., Webster, G. D. <a target="_blank" href="http://onlinelibrary.wiley.com/doi/10.1002/nau.1930110102/abstract">The History of Urodynamics.</a> Neurourol Urodyn. 11 (1), 1-21 (1992).</li>
<li>Fry, C. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Animal+models+and+their+use+in+understanding+lower+urinary+tract+dysfunction%5BTitle%5D)+AND+%22Neurourol+Urodyn%22%5BJournal%5D)">Animal models and their use in understanding lower urinary tract dysfunction.</a> Neurourol Urodyn. 29 (4), 603-608 (2010).</li>
<li>Maggi, C. A., Santicioli, P., Meli, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+nonstop+transvesical+cystometrogram+in+urethane-anesthetized+rats%3A+a+simple+procedure+for+quantitative+studies+on+the+various+phases+of+urinary+bladder+voiding+cycle%5BTitle%5D)+AND+%22J+Pharmacol+Methods%22%5BJournal%5D)">The nonstop transvesical cystometrogram in urethane-anesthetized rats: a simple procedure for quantitative studies on the various phases of urinary bladder voiding cycle.</a> J Pharmacol Methods. 15 (2), 157-167 (1986).</li>
<li>Malmgren, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cystometrical+evaluation+of+bladder+instability+in+rats+with+infravesical+outflow+obstruction%5BTitle%5D)+AND+%22J+Urol%22%5BJournal%5D)">Cystometrical evaluation of bladder instability in rats with infravesical outflow obstruction.</a> J Urol. 137 (6), 1291-1294 (1987).</li>
<li>Pandita, R. K., Fujiwara, M., Alm, P., Andersson, K. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cystometric+evaluation+of+bladder+function+in+non-anesthetized+mice+with+and+without+bladder+outlet+obstruction%5BTitle%5D)+AND+%22J+Urol%22%5BJournal%5D)">Cystometric evaluation of bladder function in non-anesthetized mice with and without bladder outlet obstruction.</a> J Urol. 164 (4), 1385-1389 (2000).</li>
<li>Chang, H. Y., Havton, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Differential+effects+of+urethane+and+isoflurane+on+external+urethral+sphincter+electromyography+and+cystometry+in+rats%5BTitle%5D)+AND+%22Am+J+Physiol+Renal+Physiol%22%5BJournal%5D)">Differential effects of urethane and isoflurane on external urethral sphincter electromyography and cystometry in rats.</a> Am J Physiol Renal Physiol. 295 (4), F1248-F1253 (2008).</li>
<li>Matsuura, S., Downie, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Effect+of+anesthetics+on+reflex+micturition+in+the+chronic+cannula-implanted+rat%5BTitle%5D)+AND+%22Neurourol+Urodyn%22%5BJournal%5D)">Effect of anesthetics on reflex micturition in the chronic cannula-implanted rat.</a> Neurourol Urodyn. 19 (1), 87-99 (2000).</li>
<li>DePaul, M. A., Lin, C. Y., Silver, J., Lee, Y. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Peripheral+Nerve+Transplantation+Combined+with+Acidic+Fibroblast+Growth+Factor+and+Chondroitinase+Induces+Regeneration+and+Improves+Urinary+Function+in+Complete+Spinal+Cord+Transected+Adult+Mice%5BTitle%5D)+AND+%22PLoS+One%22%5BJournal%5D)">Peripheral Nerve Transplantation Combined with Acidic Fibroblast Growth Factor and Chondroitinase Induces Regeneration and Improves Urinary Function in Complete Spinal Cord Transected Adult Mice.</a> PLoS One. 10 (10), e0139335(2015).</li>
<li>Kadekawa, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Characterization+of+bladder+and+external+urethral+activity+in+mice+with+or+without+spinal+cord+injury--a+comparison+study+with+rats%5BTitle%5D)+AND+%22Am+J+Physiol+Regul+Integr+Comp+Physiol%22%5BJournal%5D)">Characterization of bladder and external urethral activity in mice with or without spinal cord injury--a comparison study with rats.</a> Am J Physiol Regul Integr Comp Physiol. 310 (8), R752-R758 (2016).</li>
<li>Uvin, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+use+of+cystometry+in+small+rodents%3A+a+study+of+bladder+chemosensation%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">The use of cystometry in small rodents: a study of bladder chemosensation.</a> J Vis Exp. (66), (2012).</li>
<li>Andersson, K. E., Soler, R., Fullhase, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Rodent+models+for+urodynamic+investigation%5BTitle%5D)+AND+%22Neurourol+Urodyn%22%5BJournal%5D)">Rodent models for urodynamic investigation.</a> Neurourol Urodyn. 30 (5), 636-646 (2011).</li>
<li>Smith, P. P., Kuchel, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Continuous+uroflow+cystometry+in+the+urethane-anesthetized+mouse%5BTitle%5D)+AND+%22Neurourol+Urodyn%22%5BJournal%5D)">Continuous uroflow cystometry in the urethane-anesthetized mouse.</a> Neurourol Urodyn. 29 (7), 1344-1349 (2010).</li>
<li>Aizawa, N., Homma, Y., Igawa, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Influence+of+High+Fat+Diet+Feeding+for+20+Weeks+on+Lower+Urinary+Tract+Function+in+Mice%5BTitle%5D)+AND+%22Low+Urin+Tract+Symptoms%22%5BJournal%5D)">Influence of High Fat Diet Feeding for 20 Weeks on Lower Urinary Tract Function in Mice.</a> Low Urin Tract Symptoms. 5 (2), 101-108 (2013).</li>
<li>Bjorling, D. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Evaluation+of+voiding+assays+in+mice%3A+impact+of+genetic+strains+and+sex%5BTitle%5D)+AND+%22Am+J+Physiol+Renal+Physiol%22%5BJournal%5D)">Evaluation of voiding assays in mice: impact of genetic strains and sex.</a> Am J Physiol Renal Physiol. 308 (12), F1369-F1378 (2015).</li>
<li>Morikawa, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Effects+of+various+drugs+on+bladder+function+in+conscious+rats%5BTitle%5D)+AND+%22Jpn+J+Pharmacol%22%5BJournal%5D)">Effects of various drugs on bladder function in conscious rats.</a> Jpn J Pharmacol. 50 (4), 369-376 (1989).</li>
<li>Yaksh, T. L., Durant, P. A., Brent, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Micturition+in+rats%3A+a+chronic+model+for+study+of+bladder+function+and+effect+of+anesthetics%5BTitle%5D)+AND+%22Am+J+Physiol%22%5BJournal%5D)">Micturition in rats: a chronic model for study of bladder function and effect of anesthetics.</a> Am J Physiol. 251 (6 Pt 2), R1177-R1185 (1986).</li>
<li>Cornelissen, L. L., Misajet, B., Brooks, D. P., Hicks, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Influence+of+genetic+background+and+gender+on+bladder+function+in+the+mouse%5BTitle%5D)+AND+%22Auton+Neurosci%22%5BJournal%5D)">Influence of genetic background and gender on bladder function in the mouse.</a> Auton Neurosci. 140 (1-2), 53-58 (2008).</li>
<li>Lemack, G. E., Zimmern, P. E., Vazquez, D., Connell, J. D., Lin, V. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Altered+response+to+partial+bladder+outlet+obstruction+in+mice+lacking+inducible+nitric+oxide+synthase%5BTitle%5D)+AND+%22J+Urol%22%5BJournal%5D)">Altered response to partial bladder outlet obstruction in mice lacking inducible nitric oxide synthase.</a> J Urol. 163 (6), 1981-1987 (2000).</li>
</ol>

The authors have nothing to disclose.

Optimal material and size of intravesical tubing
To determine the effect tubing diameter has on pressure recordings, we tested different microfluidic tubes; PE50 (0.58 mm ID), polyurethane PU027 (0.4 mm ID), PE25 (0.46 mm ID), and PE10 (0.28 mm ID). For each tube, pressure was recorded with the infusion pump running at 1 mL/h, while rapidly moving the tube vertically from 0 to 30 cm. Initial in vivo experiments attempted to use PE50 tubing, but were unsuccessful due to the size of tubing compared to the mouse bladder (Figure 4A). While this supports the finding by Smith and Kuchel, who suggest that using PE50 tubing for awake cystometry in a mouse creates artifacts, making it difficult to interpret the data,12 it is important to note that others have successfully used PE50 tubing in both awake nonrestrained, awake restrained, and anesthetized mouse cystometry.9,13,14 Compared to PE50, PE10 is more flexible, which reduces the amount of tension the tube applies to the bladder as the mouse moves, reducing movement artifacts. It is important that the section of PE10 is as short as possible (&#8804; 7cm). Longer PE10 tubing leads to a greater likelihood that the pressure reading will be dampened. Although there is a theoretical advantage to using a softer, more bio-inert material like PU for intravesical implantation to decrease the inflammatory reaction, it did not result in a significant difference in post-operative bladder swelling. Further, trials which used the softer PU tubing, were associated with kinking and plugging.
Effects of surgery on bladder wall contractility and swelling
Until now, data on bladder function and bladder wall swelling, post implant of an intravesical tube, were only available for rats. According to previous studies, micturition volume was lower and micturition frequency was higher on post-operative days 1 - 3.15 It was also shown that alterations in bladder function of a rat were associated with severe swelling of the bladder, with the edema starting to subside after 3 days.16 To gain a better understanding of the change in urinary bladder function, bladder wall swelling, and repair timeline of male, C57BL/6 mice, which occur from implanting PE10 tubing, 12-hour behavioral voiding frequency was assessed using the filter paper recording method. Following the last recording, the mouse was anesthetized and the bladder was grossly assessed, harvested, fixed, and evaluated histologically. On post-surgery day 1 and 2, voiding frequency decreased and spotting increased, followed by an increase in voiding by day 3. Voiding behavior normalized by day 5. Gross evaluation of bladders at harvest and following H&#38;E staining revealed the greatest amount of suburothelial swelling on post-operative day 2 and 3, with bladders becoming similar to the control bladders at day 5 and 7 following the tube implant.
Similar to clinical urodynamic studies, the majority of laboratories used room temperature 0.9% NaCl.11 The rate of infusion in previous studies varies significantly from 10 &#181;L/min to 100 &#181;L/min.17,18 A study comparing the effects of different infusion rates on bladder function in a mouse have not been done, however, data obtained in larger animal studies recommended that slower filling rates should be used. Due to the small volume of the mouse bladder, peristaltic pumps are not appropriate and a continuous mechanism infusion pump is necessary.
The most accurate and usable FC recordings balance good transmittance of changes in pressure with limited artifacts. The results obtained using a short segment of PE10 tubing connected directly to PE50 provided an accurate measurement of pressure in the mouse urinary bladder. Pressure fluctuations caused by animal movement can be limited by anchoring the tube to the skin at the point where it exits at the nape of the neck (Figures 12 and 13). This could be achieved with the use of glue bubbles and a special anchor consisting of a metal fabric-covered plate placed subcutaneously, and the external piece, which is attached to the metal spring covering the tube. Additional methods for preventing the tube from pulling on the bladder include creating a curved subcutaneous track for the internal portion of the PE10 tubing, which provides slack in the tube, and using a swivel and tether, which prevent twisting and kinking. Based on the literature and evidence provided by these studies, the following steps are recommended to provide the most reproducible and physiologically accurate method for in vivo recording of intravesical pressure in a mouse. Use an operating microscope and microsurgical tools to implant the catheter into the dome of the urinary bladder. Allow a 5 day recovery period between the surgery and recording. Acclimate the animal in the same cage that the recording will be performed in and provide free access to food and water. Perform the experiment in a quiet environment with minimal human contact, ideally use remote video monitoring to observe the animal&#39;s behavior.

Filling cystometry (FC) is a diagnostic method that involves placing a catheter into the urinary bladder to record pressure during slow bladder filling. First introduced in 1927 as a clinical diagnostic method to evaluate lower urinary tract function, it has remained widely used.1 In research applications, FC can be used to test bladder function in healthy and diseased animal models and to study the effects of pharmacological agents. Rodent animal models are commonly used to investigate lower urinary tract function.2 In this group of mammals, FC was first developed for use in rats.3 Here, the methodology to implant a tube into the urinary bladder and perform FC has been well described and used by many researchers with an acceptable level of reproducibility.4 The availability of transgenic and knock out strains make mice a valuable species for numerous research areas, including the field of lower urinary tract dysfunction. The methodology used for performing mouse cystometry varies appreciably between laboratories, making it difficult to compare results.5
Compared to ex vivo models, FC preserves lower urinary tract anatomy, allowing the coordinated function between the bladder and its outlet during the storage and voiding phases of the micturition cycle to be assessed. Previous research shows that numerous, commonly used anesthetics suppress micturition contraction. Agents that preserve urinary bladder smooth muscle contraction (urethane, &#945;-chloralose, ketamine and xylazine), allowing the animal to micturate, still significantly reduce functional bladder capacity and suppress neurotransmission.6,7,8,9 Although technically more challenging, FC performed in awake ambulating animals preserves the functional integrity of the micturition reflex.
Lower urinary tract function is influenced by multiple factors, including post-operative bladder wall swelling, stress due to pain and discomfort, and environmental influences. Using a surgical technique that minimizes tissue damage during tube implantation and recording methods that reduce tube movement, while simultaneously allowing the animal to ambulate freely, are essential for obtaining accurate and reproducible recordings.
If performed adequately, in vivo FC in freely moving animals can provide data that reliably reflects physiological bladder function.10 FC in freely moving animals can provide data on the following parameters; Basal or baseline pressure: Minimum pressure between two micturitions. Intermicturition pressure: Mean pressure between two micturitions. Threshold pressure: Intravesical pressure immediately before micturition. Maximum pressure: Maximum bladder pressure during a micturition cycle. Spontaneous activity (or mean intermicturition oscillatory pressure): Intermicturition pressure minus basal pressure. Non-voiding contractions: Increase in intravesical pressure during the filling phase, not associated with the release of fluid. Bladder compliance: Bladder capacity divided by threshold pressure minus basal pressure. Micturition frequency: Number of micturitions per unit of time. Intermicturition interval: Period between two maximum voiding pressures. Bladder capacity: Infused volume divided by the number of micturitions. A detailed description of these parameters and standardized terminology has been previously published.11
FC can be performed using a continuous or single cycle intravesical infusion method. Continuous cystometry allows for recording of multiple micturition cycles and selecting representative data based on reproducibility. Its accuracy in measuring bladder capacity is limited due to the unknown residual volume. In addition, it is challenging to collect small voided volumes (which based on strain and sex vary between 30 and 184 &#181;L) in freely ambulating mice. Using this method to record voided volume is less accurate compared to an anesthetized preparation, but it is superior in that it avoids the suppressive effects of anesthetics on bladder function. Single cycle cystometry should be used to assess bladder capacity. In this method, the bladder is emptied by aspiration prior to infusion and capacity is calculated as a function of infusion rate multiplied by time to maximum pressure.
Although the technique of performing cystometry in small rodents has been published, it described the surgery performed in a rat and recommended that mouse cystometry should be performed under urethane anesthesia.10 The goal of this communication is to describe both the microsurgical techniques used to implant an intravesical tube into the dome of the urinary bladder and the experimental set-up used to record lower urinary tract function, in vivo, during continuous bladder filling and micturition in an awake freely moving mouse. In addition, experiments were performed to address how tubing length, diameter, and material, as well as the methodology for performing in vivo FC, affect the recording. This experimental protocol summarizes previously published techniques and proposes a number of modifications based on experimental results.

<table><tbody><tr><td>Polyethylene (PE) 10 tubing</td><td>Instech</td><td>BTPE-10</td><td>Fits 30G connectors/plugs</td></tr><tr><td>Polyethylene (PE) 50 tubing</td><td>Instech</td><td>BTPE-50</td><td>Fits 22G connectors/plugs</td></tr><tr><td>22 G single channel stainless steel swivel</td><td>Instech</td><td>375/22</td><td></td></tr><tr><td>High Carbon Steel Utility Extension Spring (9/64&quot; OD)</td><td>Grainger</td><td>1NAH1</td><td>Protects PE50 tubing - Cut to length</td></tr><tr><td>22 G connector</td><td>Instech</td><td>SP22/12</td><td></td></tr><tr><td>Yutaoz Professional Hot Melt Adhesive Glue Gun</td><td>Yutaoz</td><td></td><td>Use low temperature setting (100 &amp;deg;C) - Any hot melt glue gun with an adjustable temperature range will work</td></tr><tr><td>Surebonder DT-2010 all purpose glue stick</td><td>Surebonder</td><td></td><td>Any all purpose hot glue will work</td></tr><tr><td>Dumont #5 curved microforceps</td><td>World Precision Instruments</td><td>500232</td><td></td></tr><tr><td>Dumont #7 curved microforceps</td><td>World Precision Instruments</td><td>14188</td><td></td></tr><tr><td>Mini dissecting scissors - straight</td><td>World Precision Instruments</td><td>503240</td><td></td></tr><tr><td>Micro mosquito forceps (12.5 cm)</td><td>World Precision Instruments</td><td>500451</td><td></td></tr><tr><td>Dissecting scissors - straight</td><td>World Precision Instruments</td><td>14393</td><td></td></tr><tr><td>Castroviejo Needle Holder</td><td>World Precision Instruments</td><td>503258</td><td></td></tr><tr><td>Isoflurane, USP</td><td>Phoenix</td><td></td><td>2%, 1 L/min flow rate</td></tr><tr><td>Buprenorphine</td><td></td><td></td><td>0.05 mg/kg</td></tr><tr><td>0.9% Sodium Chloride Irrigation, USP</td><td>Baxter</td><td></td><td></td></tr><tr><td>6-0 Ethilon black monofilament, non-absorbable suture</td><td>Ethicon</td><td></td><td>Bladder tie</td></tr><tr><td>6-0 Vicryl violet braided, absorbable suture</td><td>Ethicon</td><td></td><td>Muscle suture, running</td></tr><tr><td>6-0 Prolene blue monofilament, non-absorbable suture</td><td>Ethicon</td><td></td><td>Skin suture, vertical mattress, buried interrupted</td></tr><tr><td>KD Legato 210 infuse/withdraw pump</td><td>KD Scientific</td><td></td><td>1.5 mL/hr</td></tr><tr><td>Disposable pressure transducer</td><td>Digitimer</td><td>NL108T2</td><td></td></tr><tr><td>Pressure Amplifier</td><td>Digitimer</td><td>NL108A</td><td></td></tr><tr><td>Power1401-3 data acquisition interface</td><td>Digitimer</td><td></td><td></td></tr><tr><td>Spike2&amp;nbsp;</td><td>Cambridge Electronic Design Limited</td><td></td><td>PC pressure recording software</td></tr><tr><td>Leica MZ6 surgical operating microscope (3.2 - 20X)</td><td>Leica Microsystems</td><td></td><td>Magnification</td></tr></tbody></table>

evaluating the procedure for performing awake cystometry in a mouse model

Awake filling cystometry has been used for a long time to evaluate bladder function in freely moving mice, however, the specific methods used, vary among laboratories. The goal of this study was to describe the microsurgical procedure used to implant an intravesical tube and the experimental technique for recording urinary bladder pressure in an awake, freely moving mouse. In addition, experimental data is presented to show how surgery, as well as tubing type and size, affect lower urinary tract function and recording sensitivity. The effect of tube diameter on pressure recording was assessed in both polyethylene and polyurethane tubing with different internal diameters. Subsequently, the best performing tube from both materials was surgically implanted into the dome of the urinary bladder of male C57BL/6 mice. Twelve-hour, overnight micturition frequency was recorded in healthy, intact animals and animals 2, 3, 5, and 7 days post-surgery. At harvest, bladders were assessed for signs of swelling using gross observation and were subsequently processed for pathological analysis. The greatest extent of bladder swelling was observed on day 2 and 3, which correlated with behavioral voiding data showing significantly impaired bladder function. By day 5, bladder histology and voiding frequency had normalized. Based on the literature and evidence provided by our studies, we propose the following steps for in vivo recording of intravesical pressure and voided volume in an awake mouse: 1) Perform the surgery using an operating microscope and microsurgical tools, 2) Use polyethylene-10 tubing to minimize movement artifacts, and 3) Perform cystometry on post-operative day 5, when bladder swelling resolves.

There was no significant difference between the tubing materials and diameters in the consistency of pressure rise and fall within the system during tube occlusion. Bladder wall swelling post intravesical tube implantation was significant for both polyethylene (PE) and polyurethane (PU) materials. On day 2, severe submucosal swelling developed. It occupied half the cross-section of the bladder, leading to obstruction of the lumen. On day 5, the edema resolved completely, leaving the submucosal areas infiltrated with inflammatory cells that partially invaded the muscularis. On day 7, the inflammatory infiltration was reduced significantly and the bladder wall histology returned to normal (Figure 8). The greatest extent of tissue swelling observed on day 2 and day 3 correlated with behavioral voiding data showing significantly impaired bladder function (Figure 9). Voiding frequency normalized by post-operative day 5.
Intravesical pressure in an awake freely moving mouse (with minimal movement artifacts) is characterized by a baseline pressure of 10-15 cm H2O, which may remain unchanged or gradually increase by no more than 10-cm H2O during the filling cycle, followed by a sudden, pulsatile pressure increase and then drop during voiding (Figures 10 and 11).
<img alt="Figure 1" src="/files/ftp_upload/55588/55588fig1.jpg" /> 
Figure 1: PE10 Tubing for Implantation into the Urinary Bladder. (A) A 7 cm piece of PE10 tubing with drops of hot glue at 4.5, 5, and 5.5 cm from the flared end. (B and C) A detailed picture showing the flared end of the tube (used to secure tube in bladder). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55588/55588fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/55588/55588fig2.jpg" /> 
Figure 2: Plug for the External Portion of the PE10 Bladder Tubing. Plug is made from a 30 G needle and hot glue. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55588/55588fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/55588/55588fig3.jpg" /> 
Figure 3: Course of the Bladder Tubing. Schematic line drawing illustrating the placement of the tubing through the abdomen and its subcutaneous route towards the nape of the neck. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55588/55588fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/55588/55588fig4.jpg" /> 
Figure 4: Bladder/Tubing Comparison and Steps used to Insert and Secure the Tube using a Lose Monofilament Suture. Intraoperative photographs depicting: (A) Picture comparing PE50 and PE10 to a mouse urinary bladder. (B) A small loop of 6-0 monofilament suture placed around the bladder. A pair of #5 microforceps clasps the dome of the bladder while a 21 G needle is used to make the cystotomy. (C) Without releasing the dome of the bladder, a pair of #5 microforceps in the opposite hand probes the hole before inserting the PE10 catheter. (D) PE10 catheter secured in the dome of the bladder with 6-0 monofilament suture. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55588/55588fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/55588/55588fig5.jpg" /> 
Figure 5: A Purse String Suture can be used as an Alternative Method for Securing the Tube in the Bladder. (A) Purse string suture in the dome of the bladder. (B) PE10 tubing inserted through a center of the purse string. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55588/55588fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" src="/files/ftp_upload/55588/55588fig6.jpg" /> 
Figure 6: Experimental Set-up. Syringe containing 0.9% NaCl in the infusion pump connected in series to the pressure transducer and intravesical catheter. The computer screen in the bottom right shows three reproducible micturition cycles. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55588/55588fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" src="/files/ftp_upload/55588/55588fig7.jpg" /> 
Figure 7: Experimental Recording Set-up. (A) 22 G swivel suspended over a recording cage and balance. (B) Photograph showing entire length of the external portion of the infusion tubing with a 22 G swivel and a PE50 tether. (C) A 22 G swivel with a spring sheath covering PE50 tubing. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55588/55588fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" src="/files/ftp_upload/55588/55588fig8.jpg" /> 
Figure 8: Histological Evaluation of the Response of the Urinary Bladder Wall to an Implanted PE10 Tube. Cross-sections of the urinary bladder stained with hematoxylin and eosin (H&#38;E) before, 2, 3, 5 and 7 days post surgery. Bladder wall swelling resolved on post-operative day 5. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55588/55588fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" src="/files/ftp_upload/55588/55588fig9.jpg" /> 
Figure 9: Functional Bladder Evaluation using Voiding Spot Assay. Urine spots on filter paper viewed with UV light documenting the representative micturition pattern on day 0 (prior to tube implant), 2, 3, 4, 5 and 7 post-intravesical tube implant. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55588/55588fig9large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 10" src="/files/ftp_upload/55588/55588fig10.jpg" /> 
Figure 10: Cystometrogram. Representative trace of intravesical bladder pressure in an awake, freely moving mouse. Trace showing 3 reproducible micturition cycles. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55588/55588fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 11" src="/files/ftp_upload/55588/55588fig11.jpg" /> 
Figure 11: Micturition Phase. Trace depicting the micturition phase with high frequency oscillations during the initial increase in pressure, peak pressure, and a rapid pressure drop to baseline. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55588/55588fig11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 12" src="/files/ftp_upload/55588/55588fig12.jpg" /> 
Figure 12: Anchoring the Tether. (A) Anchor used to secure the PE10 catheter in the animal and prevent the tube from puling on the bladder. The disk is attached to the spring sheath. (B) Internal PE10 catheter connected to the external PE50 tubing. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55588/55588fig12large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 13" src="/files/ftp_upload/55588/55588fig13.jpg" /> 
Figure 13: Swivel Anchor. Subcutaneous portion of the anchor consisting of (1) a disc made of fabric and (2) a metal loop.

Watch this Scientific Journal Video about Evaluating the Procedure for Performing Awake Cystometry in a Mouse Model at JoVE.com

Evaluating the Procedure for Performing Awake Cystometry in a Mouse Model

This study describes the surgical procedures and experimental techniques for performing awake cystometry in a freely moving mouse. In addition, it provides experimental evidence to support its optimization and standardization.

Awake filling cystometry has been used for a long time to evaluate bladder function in freely moving mice, however, the specific methods used, vary among ...

evaluating-procedure-for-performing-awake-cystometry-mouse

Nanyang Technological University

Research

JoVE Journal

Medicine

12.6K Views.  University of Vermont. This study describes the surgical procedures and experimental techniques for performing awake cystometry in a freely moving mouse. In addition, it provides experimental evidence to support its optimization and standardization.

Video: Evaluating the Procedure for Performing Awake Cystometry in a Mouse Model

A Murine Orthotopic Bladder Tumor Model and Tumor Detection System

This protocol describes the generation of bladder tumors in female C57BL/6J mice using the murine bladder cancer cell line MB49, which has been modified to secrete human Prostate Specific Antigen (PSA), and the procedure for the confirmation of tumor implantation. In brief, mice are anesthetized using injectable drugs and are made to lay in the dorsal position. Urine is vacated from the bladder and 50 &#181;L of poly-L-lysine (PLL) is slowly instilled at a rate of 10 &#181;L/20 s using a 24 G IV catheter. It is left in the bladder for 20 min by stoppering the catheter. The catheter is removed and PLL is vacated by gentle pressure on the bladder. This is followed by instillation of the murine bladder cancer cell line (1 x 105 cells/50 &#181;L) at a rate of 10 &#181;L/20 s. The catheter is stoppered to prevent premature evacuation. After 1 h, the mice are revived with a reversal drug, and the bladder is vacated. The slow instillation rate is important, as it reduces vesico-ureteral reflux, which can cause tumors to occur in the upper urinary tract and in the kidneys. The cell line should be well re-suspended to reduce clumping of cells, as this can lead to uneven tumor sizes after implantation.
This technique induces tumors with high efficiency. Tumor growth is monitored by urinary PSA secretion. PSA marker monitoring is more reliable than ultrasound or fluorescence imaging for the detection of the presence of tumors in the bladder. Tumors in mice generally reach a maximum size that negatively impacts health by about 3 - 4 weeks if left untreated. By monitoring tumor growth, it is possible to differentiate mice that were cured from those that were not successfully implanted with tumors. With only end-point analysis, the latter may be mistakenly assumed to have been cured by therapy.

This protocol describes the generation of murine orthotopic bladder tumors in female C57BL/6J mice and the monitoring of tumor growth.

This protocol describes the generation of bladder tumors in female C57BL/6J mice using the murine bladder cancer cell line MB49, which has been modified ...

Cancer Research

Flat-floored Air-lifted Platform: A New Method for Combining Behavior with Microscopy or Electrophysiology on Awake Freely Moving Rodents

It is widely acknowledged that the use of general anesthetics can undermine the relevance of electrophysiological or microscopical data obtained from a living animal&#8217;s brain. Moreover, the lengthy recovery from anesthesia limits the frequency of repeated recording/imaging episodes in longitudinal studies. Hence, new methods that would allow stable recordings from non-anesthetized behaving mice are expected to advance the fields of cellular and cognitive neurosciences. Existing solutions range from mere physical restraint to more sophisticated approaches, such as linear and spherical treadmills used in combination with computer-generated virtual reality. Here, a novel method is described where a head-fixed mouse can move around an air-lifted mobile homecage and explore its environment under stress-free conditions. This method allows researchers to perform behavioral tests (e.g., learning, habituation or novel object recognition) simultaneously with two-photon microscopic imaging and/or patch-clamp recordings, all combined in a single experiment. This video-article describes the use of the awake animal head fixation device (mobile homecage), demonstrates the procedures of animal habituation, and exemplifies a number of possible applications of the method.

This method creates a tangible, familiar environment for the mouse to navigate and explore during microscopic imaging or single-cell electrophysiological recordings, which require firm fixation of the animal&#8217;s head.

It is widely acknowledged that the use of general anesthetics can undermine the relevance of electrophysiological or microscopical data obtained from a ...

Behavior

Evaluation of Biomaterials for Bladder Augmentation using Cystometric Analyses in Various Rodent Models

Renal function and continence of urine are critically dependent on the proper function of the urinary bladder, which stores urine at low pressure and expels it with a precisely orchestrated contraction. A number of congenital and acquired urological anomalies including posterior urethral valves, benign prostatic hyperplasia, and neurogenic bladder secondary to spina bifida/spinal cord injury can result in pathologic tissue remodeling leading to impaired compliance and reduced capacity1. Functional or anatomical obstruction of the urinary tract is frequently associated with these conditions, and can lead to urinary incontinence and kidney damage from increased storage and voiding pressures2. Surgical implantation of gastrointestinal segments to expand organ capacity and reduce intravesical pressures represents the primary surgical treatment option for these disorders when medical management fails3. However, this approach is hampered by the limitation of available donor tissue, and is associated with significant complications including chronic urinary tract infection, metabolic perturbation, urinary stone formation, and secondary malignancy4,5.
Current research in bladder tissue engineering is heavily focused on identifying biomaterial configurations which can support regeneration of tissues at defect sites. Conventional 3-D scaffolds derived from natural and synthetic polymers such as small intestinal submucosa and poly-glycolic acid have shown some short-term success in supporting urothelial and smooth muscle regeneration as well as facilitating increased organ storage capacity in both animal models and in the clinic6,7. However, deficiencies in scaffold mechanical integrity and biocompatibility often result in deleterious fibrosis8, graft contracture9, and calcification10, thus increasing the risk of implant failure and need for secondary surgical procedures. In addition, restoration of normal voiding characteristics utilizing standard biomaterial constructs for augmentation cystoplasty has yet to be achieved, and therefore research and development of novel matrices which can fulfill this role is needed.
In order to successfully develop and evaluate optimal biomaterials for clinical bladder augmentation, efficacy research must first be performed in standardized animal models using detailed surgical methods and functional outcome assessments. We have previously reported the use of a bladder augmentation model in mice to determine the potential of silk fibroin-based scaffolds to mediate tissue regeneration and functional voiding characteristics.11,12 Cystometric analyses of this model have shown that variations in structural and mechanical implant properties can influence the resulting urodynamic features of the tissue engineered bladders11,12. Positive correlations between the degree of matrix-mediated tissue regeneration determined histologically and functional compliance and capacity evaluated by cystometry were demonstrated in this model11,12. These results therefore suggest that functional evaluations of biomaterial configurations in rodent bladder augmentation systems may be a useful format for assessing scaffold properties and establishing in vivo feasibility prior to large animal studies and clinical deployment. In the current study, we will present various surgical stages of bladder augmentation in both mice and rats using silk scaffolds and demonstrate techniques for awake and anesthetized cystometry.

Surgical stages of bladder augmentation are described using 3-D scaffolds in murine and rat models. To test the efficacy of biomaterial configurations for use in bladder augmentation, techniques for both awake and anesthetized cystometry are presented.

Renal function and continence of urine are critically dependent on the proper function of the urinary bladder, which stores urine at low pressure and ...

Bioengineering

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 ...

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.

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 ...

In Vivo Luminal Measurement of Distension-Evoked Urothelial ATP Release in Rodents

ATP, released from the urothelium in response to bladder distension, is thought to play a significant sensory role in the control of micturition. Therefore, accurate measurement of urothelial ATP release in a physiological setting is an important first step in studying the mechanisms that control purinergic signaling in the urinary bladder. Existing techniques to study mechanically evoked urothelial ATP release utilize cultured cells plated on flexible supports or bladder tissue pinned into Ussing chambers; however, each of these techniques does not fully emulate conditions in the intact bladder. Therefore, an experimental setup was developed to directly measure ATP concentrations in the lumen of the rodent urinary bladder.
In this setup, the bladders of anesthetized rodents are perfused through catheters in both the dome of the bladder and via the external urethral orifice. Pressure in the bladder is increased by capping the urethral catheter while perfusing sterile fluid into the bladder through the dome. Measurement of intravesical pressure is achieved using a pressure transducer attached to the bladder dome catheter, akin to the setup used for cystometry. Once the desired pressure is reached, the urethral catheter's cap is removed, and fluid collected for ATP quantification by luciferin-luciferase assay. Through this experimental setup, the mechanisms controlling both mechanical and chemical stimulation of urothelial ATP release can be interrogated by including various agonists or antagonists into the perfusate or by comparing results between wildtype and genetically modified animals.

In Vivo Luminal Measurement of Distension-Evoked Urothelial ATP Release in Rodents

This protocol describes the procedure for measuring ATP concentrations in the lumen of the bladder in an anesthetized rodent.

ATP, released from the urothelium in response to bladder distension, is thought to play a significant sensory role in the control of micturition. ...

Cystometric and External Urethral Sphincter Measurements in Awake Rats with Implanted Catheter and Electrodes Allowing for Repeated Measurements

Lower urinary tract function is mainly assessed by means of cystometric bladder function analysis in rodents. Conventional cystometries are usually performed as terminal analysis under urethane anesthesia. It is well known that anesthetic drugs can influence bladder function. Hence, the aim of this technique is to perform cystometric measurements of the urinary bladder and external urethral sphincter in lightly restrained awake rats. For this purpose, a bladder catheter is implanted into the bladder dome. Subsequently, two electrodes are implanted bilateral to the external urethral sphincter and a ground electrode is sutured to a non-responsive skeletal muscle. The bladder catheter and the three electrodes are finally tunneled subcutaneously to the neck region and affixed to a harness. With this technique, the lower urinary tract can be measured at multiple time points in the same animal to assess lower urinary tract function. The main application of this technique is the follow-up of simultaneous urinary bladder and external urethral sphincter function in awake healthy rats and after induction of a disease or injury. Moreover, subsequent lower urinary tract monitoring can be performed during evaluation of the disease/injury and to monitor treatment efficacy.

This protocol first describes the surgical procedure of the permanent implantation of a urinary bladder catheter combined with external urethral sphincter electrodes, and second, the measurement of the function of the urinary bladder and external urethral sphincter in implanted awake animals.

Lower urinary tract function is mainly assessed by means of cystometric bladder function analysis in rodents. Conventional cystometries are usually ...

Biology

Precise Urodynamic Measurement in Awake Mice Using Cyanoacrylate Skin Barriers

Optimizing Mouse Urodynamic Techniques for Improved Accuracy

Accurate measurement of urinary parameters in awake mice is crucial for understanding lower urinary tract (LUT) dysfunction, particularly in conditions like neurogenic bladder post-traumatic spinal cord injury (SCI). However, conducting cystometry recordings in mice presents notable challenges. When mice are in a prone and restricted position during recording sessions, urine tends to be absorbed by the fur and skin, leading to an underestimation of voided volume (VV). The goal of this study was to enhance the accuracy of cystometry and external urethral sphincter electromyography (EUS-EMG) recordings in awake mice. We developed a unique method utilizing cyanoacrylate adhesive to create a waterproof skin barrier around the urethral meatus and abdomen, preventing urine absorption and ensuring precise measurements. Results show that after applying the cyanoacrylate, the sum of VV and RV remained consistent with the infused saline volume, and there were no wet areas observed post-experiment, indicating successful prevention of urine absorption. Additionally, the method simultaneously stabilized the electrodes connected with the external urethral sphincter (EUS), ensured stable electromyography (EMG) signals, and minimized artifacts caused by the movement of the awakened mouse and manipulation of the experimenter. Methodological details, results, and implications are discussed, highlighting the importance of improving urodynamic techniques in preclinical research.

Author Spotlight: Enhanced Urodynamic Method for Precise Urine Measurement in Awake Mice with Neurogenic Bladder

This protocol provides a guide for waterproofing the skin with cyanoacrylate to prevent urine absorption by fur and skin. It includes instructions for applying the glue to the skin, implanting a bladder catheter, and electrodes for cystometry and external urethral sphincter electromyography recordings in awake mice.

Accurate measurement of urinary parameters in awake mice is crucial for understanding lower urinary tract (LUT) dysfunction, particularly in conditions ...

Neuroscience

Modeling Orthotopic Bladder Tumor in Mouse Model: A Procedure for Intravesical Administration of Cancer Cells into Murine Bladder

Source: Tham, S. M. et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/55078" target="_blank">A Murine Orthotopic Bladder Tumor Model and Tumor Detection System.</a>&#160;J. Vis. Exp.&#160;(2017)
In this video, we describe the intravesical procedure of delivering cancer cells into the bladder of a mouse model. This procedure helps generate an orthotopic bladder tumor model that can be used to develop therapeutic strategies.

Modelowanie ortotopowego guza pęcherza moczowego w modelu mysim: procedura dopęcherzowego podawania komórek rakowych do mysiego pęcherza moczowego

Source: Tham, S. M. et al. A Murine Orthotopic Bladder Tumor Model and Tumor Detection System.&#160;J. Vis. Exp.&#160;(2017)
In this video, we describe ...

JoVE Encyclopedia of Experiments

Urinary Tract Cancer

In vivo studies

Urine Depletion and Bladder Rinsing in Mouse Model: A Catheter-based Procedure to Empty and Wash Murine Urinary Bladder

Source: Yu, C. et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/57649" target="_blank">Novel Method of Plasmid DNA Delivery to Mouse Bladder Urothelium by Electroporation.</a> J. Vis. Exp. (2018)
In this video, we describe a catheter-based procedure to empty and rinse the urinary bladder of a mouse model. The prepped mouse can then be used for further experimental procedures.

Wyczerpanie moczu i płukanie pęcherza moczowego w modelu mysim: procedura oparta na cewniku do opróżniania i mycia mysiego pęcherza moczowego

Source: Yu, C. et al. Novel Method of Plasmid DNA Delivery to Mouse Bladder Urothelium by Electroporation. J. Vis. Exp. (2018)
In this video, we describe ...

Evaluating the Procedure for Performing Awake Cystometry in a Mouse Model

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