Name: Evaluating the Procedure for Performing Awake Cystometry in a Mouse Model (Video) | JoVE Journal
Uploaded: 2017-05-20T13:00:00.000+00:00
Duration: 9 min 31 s
Description: Learn awake cystometry in mice using microsurgical intravesical tube implantation. This standardized method enables reproducible bladder function assessment in conscious animals for urological research.

ai_agent_IP

ai_agent

maintenance

instantTrialDurationMinutes

isBiopharma

enableNewMyJoveSectionDesign

educationFeatures

seoMetaApi

ai_copilot_institution_blacklist

ai_chat_institution_blacklist

freeAccessWindow

ssoRoleCapture

scheduledMaintenanceBanner

joveLabsFeatures

disable_autoplay_for_bots

extended_trial_duration

5 minutes

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...

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

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 submu...

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 ...

The overall goal of this microsurgical procedure is to implant an intravesical tube into the urinary bladder and to record the lower urinary tract function in an awake mouse. This method can help in the acquisition of reproducible experimental data from awake cystometry and the testing of bladder function in healthy and diseased mouse models. The main goal of this technique is to provide a standardized method based on experimental data for performing filling cystometry in a mouse model.To prepare the tube for implantation, first cut a seven centimeter piece of pe10 tubing, and slowly advance one end toward an open flame to create a flare, quickly withdrawing the tube as soon as the flare develops. Then use the low heat setting on a glue gun to apply three drops of all purpose hot glue at 4.5, 5, and 5.5 centimeters from the flared end of the tube. Use scissors to trim irregularities from the glue bubbles.Apply hot glue to one end of a 30 gauge needle, which will be used to plug the pe10 tubing after it is implanted. Next confirm the lack of response to toe pinch and apply ointment to the animal's eyes. After sterilizing the upper back and lower abdomen with alcohol and betadine, make a 1.5 centimeter incision between the scalpula, and place the animal in the supine position on top of a 37 degree celsius heating pad covered with sterile drapes under a dissecting microscope.Now make a 1.5 centimeter lower midline abdominal skin incision. Followed by a matching incision through the fascia along the linea alba and muscle to expose the dome and upper half of the urinary bladder. Keeping the abdominal viscera moist with warm physiological saline, rotate the animal onto it's side to access the incision on the nape of the neck.Push a narrow hemostat subcutaneously through the incision to start a subcutaneous channel on the back and continuing along the side of the animal. Once the tip of the instrument reaches the bottom of the ribcage, turn the tip towards the midline in the inside of the abdomen and continue advancing the hemostat until the tip is exposed at the abdominal incision under the muscular layer. Once the hemostat tip is inside the incision, pinch the skin above the shoulder and apply traction to help stabilize the animal until the instrument tip is seen at the abdominal incision.Now use the hemostat to grasp the non flared end of the tubing and slowly retract the hemostat, pulling the end of the tube out of the incision at the back of the neck and adjusting the flared end of the tubing so that it lies directly above the dome of the bladder. Stabilize the bladder with a small roll of lint free tissue. Place a loosely tied, not absorbable 6-O monofilament suture on top of the bladder dome.Using the non dominant hand and the Dumont number seven curved micro forceps, grasp the dome of the bladder and use a 21 gauge needle in the dominant hand to make a cystotomy in the dome apex. Gently probe the cystotomy with a closed pair of number five curved micro forceps to make sure the catheter can easily pass through the hole and place the flared end of the pe10 catheter into the bladder, pushing the flare down to the bladder neck so that it does not slip out while being secured. With the monofilament tie placed anterior to the tubing, tighten the 6-O suture around the dome of the bladder and tubing as high up on the bladder as possible to avoid artificially reducing the bladder capacity.Alternatively, place a loose purse-string suture on the dome of the bladder, make a cystotomy as just demonstrated, and insert the flared end of the catheter into the puncture. Then tie the suture around the dome and the tubing as demonstrated for the flat suture. In either case take care to secure the tubing without causing extensive injury to the bladder wall.If too much tissue is taken, the bladder capacity will be reduced. If too little is taken, the connection will leak. To test the seal and position of the tube in the bladder, attach a 0.5 milliliter insulin syringe equipped with a 30 gauge needle to the distal end of the tube and slowly fill the bladder with 0.1 to 0.2 milliliters of 0.9%sodium chloride until a drop appears at the urethral orifice.If the seal is incomplete, loop suture tails around the bladder and retie. Then empty the bladder by aspiration. If no leaks occur at the dome, brace the bladder with a pair of curved micro forceps and gently pull on the tubing until the flare is resting against the inside of the bladder dome.Retest the seal using the previously described method. Trim the tails of the suture, remove the tissue roll, and return the bladder to it's normal position. Then use a running 6-O suture to close the abdominal wall in two layers.And gently rotate the animal onto it's abdomen. Insert the subcutaneous portion of the metal anchor into the intrascopular incision, and use a 6-O monofilament to secure the tube and anchor with a vertical mattress suture. Confirm that a glue bubble remains above and below the skin to prevent the tube from pulling out and cut the tube approximately two centimeters above the skin.Then gently insert a 30 gauge plug into the end of the tube to prevent urine from leaking out. Before performing the reading, use pe50 tubing to connect the infusion pump, pressure transducer, and 22 gauge swivel. Then place the anesthetized experimental animal in the prone position and remove the 30 gauge plug.Connect the tether to the anchor, slide the bladder catheter into the end of the pe50, and use hot glue to form a water tight seal. Place the mouse onto the wire floor. And begin the recording.Then when the animal recovers from the anesthesia and the bladder pressure stabilizes, begin infusing 0.9%sodium chloride at 0.6 milliliters per hour rate. Typically, on day two after catheter insertion, severe submucosal swelling that occupies half of the cross section of the bladder develops leading to obstruction of the lumen. On day five the edema resolves completely leaving the submucosal areas infiltrated with inflammatory cells that partially invaded the muscularis.The greatest extent of the tissue swelling observed on days two and day three correlates with the corresponding behavioral voiding with the animals demonstrating a significantly impaired bladder function at these time points. As expected from the histologic data, the voiding frequency normalizes by post operative day five. Intravesical pressure in an awake freely moving animal with minimal movement artifacts is characterized by a baseline pressure of 10 to 15 centimeters of water which may remain unchanged or may gradually increase by no more than 10 centimeters of water during the filling cycle.Followed by a sudden pulse style pressure increase and drop during voiding. Once mastered, this technique can be completed in 30 minutes if it is performed properly. While attempting this procedure it is important to remember to minimize the damage to the bladder to preserve the physiology of the tissue.Following this procedure, other methods, like single cycle filling cystometry can be performed to answer additional questions about the functional bladder capacity. After watching this video, you should have a good understanding of how to microsurgically implant a tube into the urinary bladder and to use awake filling cystometey to evaluate lower urinary tract function in mice.

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

Research

JoVE Journal

Medicine

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

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.

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

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.

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

This video demonstrates a technique to assess the anti-tumor activity of Helicobacter pylori neutrophil-activating protein, or HP-NAP, in a mouse bladder cancer model. HP-NAP interacts with tumor-cell neutrophils to produce pro-inflammatory cytokines, inducing the differentiation of naïve T cells into type 1 T helper (Th1) and type 1 cytotoxic (Tc1) T cell phenotypes. The Th1 and Tc1 cells secrete interferon-gamma, inhibiting tumor angiogenesis and promoting cytotoxic T cell-mediated tumor...

Evaluating the Anti-Tumor Efficacy of a Bacterial Immunomodulatory Protein in a Murine Model

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.

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

The ability to model urinary tract infections (UTI) is crucial in order to be able to understand bacterial pathogenesis and spawn the development of novel therapeutics. This work’s goal is to demonstrate mouse models of experimental UTI and catheter associated UTI that recapitulate and predict findings seen in humans.

Establishment and Characterization of UTI and CAUTI in a Mouse Model

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

A Murine Orthotopic Bladder Tumor Model and Tumor Detection System

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.

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

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

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

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.

Optimizing Mouse Urodynamic Techniques for Improved Accuracy

Learn awake cystometry in mice using microsurgical intravesical tube implantation. This standardized method enables reproducible bladder function assessment in conscious animals for urological research.

Evaluating the Procedure for Performing Awake Cystometry in a Mouse Model

Explore More Videos