This study was supported by NIH-NINDS (R21NS130241), IND DEPT HLTH (55051, 74247, 74244), and US ARMY (HT94252310700).

The animal study protocol was approved by the Institutional Animal Care and Use Committee of Indiana University School of Medicine. Approval Code: 21098MD/R/MSS/HZ Approval Date: 29 September 2021.
1. Preparation of catheter
<ol>
	<li>Cut a 30 cm polyethylene PE-30 tube (.017 inch x .030 inch). Use a lighter to flare one end of the tube, ensuring it does not touch the flame and withdrawing the lighter once the tube has formed an appropriately round, bell-shaped tip. Any residue from the flaring procedure is then carefully cleaned or removed before proceeding with the surgery.</li>
	<li>Carefully insert about 3/4 of the 25G needle into the other end of the tube. Prepare a 1 mL syringe and fill it with sterile 0.9% NaCl. Connect the syringe to the 25 G needle.</li>
	<li>Gently infuse saline to check for a proper round tip and no leakage from the ends of the needle. Ensure that no pressure is felt and saline flows smoothly through the catheter.</li>
</ol>
2. Preparation of electrodes
<ol>
	<li>Prepare 2 steel wires of 20 cm in length. Take the steel wires and apply sand polish to both ends of the coating zone to strip 5 mm of the wire.</li>
	<li>Take a 25G needle and insert it on one side of the wire. Make sure to insert the needle carefully to avoid damaging the wire. Bend the stripped part of the wire like a hook. The hook will help to connect the wire to the EUS muscle.</li>
	<li>Use solder to attach the pin to the other end of the striped wire. Soldering will help secure the pin to the wire and ensure a strong connection. Make sure to heat the tin-lead soldering until it melts and covers the wire and pin.</li>
</ol>
3. Preparation of animal
<ol>
	<li>House female C57BL/6 mouse (8 weeks old, 18-20 g body weight) in the animal facility according to Institutional Animal Care with a 12 h light-dark cycle and unlimited access to water and standard food pellets. 
	NOTE: Prior to the experimental procedures, mice were acclimated to the restraint apparatus over a period of three days. During this time, each mouse was placed in the restraint for 10-15 minutes daily, allowing them to become familiar with the setup. Mice were monitored for signs of distress, and adjustments were made to promote a calm experience. This acclimation aimed to reduce anxiety and enhance animal welfare during the experiments</li>
</ol>
4. Anesthesia during surgery
<ol>
	<li>Place the animals in a 2% isoflurane and pure oxygen (1 L/min) chamber. Confirm the animal&#39;s complete anesthesia using negative toe-pinch examination before transferring it to the mask. Once confirmed, change the gas condition to a mask.</li>
	<li>Ensure that the anesthesia mask is fixed in the 
	appropriate position on the sterile surgical field. Place the animal supine on the heating pad covered with the sterile drape with its nose in an inhalation small mask (0.8-1 L/min with 2% isoflurane) to continue administering anesthesia.</li>
</ol>
5. Surgical preparation
<ol>
	<li>Fix the limbs of the animal with tape. Use an electric razor to shave the fur of the lower abdomen and around the urethral meatus (genital region).&#160;Administer 3 mg/kg Meloxicam (5 mg/ml) subcutaneously as a pre-emptive analgesic to the mouse before surgery to ensure effective pain management.</li>
	<li>Apply an eye ointment to prevent any potential dryness in the eyes. Prepare the shaved area using the povidone-iodine solution and wipe away the solution with 70% ethanol. Put a sterile drape on the surgery region.</li>
	<li>Autoclave all surgical tools and electrode wires&#160;at 121&#176;C for 15&#8211;20 minutes to ensure sterility prior to the procedure. Soak the polyethylene tube in a&#160;glutaraldehyde or ethylene oxide&#160;for&#160;20&#8211;30 minutes,&#160;then rinse thoroughly with sterile saline to remove any residues</li>
</ol>
6. Surgical procedure
<ol>
	<li>Bladder catheter implantation
	<ol>
		<li>Under the surgical microscope, utilizing Metzenbaum straight, blunt scissors, create a 1-2 cm incision in the midline of the abdominopelvic skin. Proceed to incise the fascia and muscles in the midline to expose the bladder through the incisional wound.</li>
		<li>Once the bladder is visible through the incisional wound, proceed to retract any surrounding organs or tissues as needed to obtain a clear view of the surgical field. Take care to avoid any unnecessary manipulation or tension on the bladder, as this can lead to complications such as urinary leakage or injury to surrounding structures.</li>
		<li>Grasp the bladder dome and place a purse string by utilizing a 5-0 nonabsorbable monofilament suture with a taper-tip needle.</li>
		<li>Using micro scissors, create a small cystostomy within the purse string and make a hole until urine flows out.</li>
		<li>Grasp the round tip end of the catheter and insert it through the hole. Once the tip of the tube has passed through the hole, suture the purse string around the tube. Then, gently pull the tube outward until the tip is felt under the suture.</li>
		<li>Slowly infuse 1 mL of saline from the other end of the tube to distend the bladder. Check for any leakage around the catheter. If leakage is present, place an additional suture.</li>
		<li>Once the saline comes out of the urethra, withdraw the saline to decompress the bladder.</li>
	</ol>
	</li>
	<li>EUS electrodes implantation (Figure 1)
	<ol>
		<li>Use surgical scissors to extend the abdominal incision up to the pelvic floor.</li>
		<li>In line with the bladder, move the muscles and membranes to the pudendal canals and locate the urethra and external sphincter muscle. Be careful not to hurt iliac and middle caudal vessels and pudendal nerves.</li>
		<li>Puncture the skin bilaterally, 1 cm away from the urethral meatus, using the needle containing the electrode.</li>
		<li>Carefully grab the hook tip with forceps and gently pull the needle away from the skin.</li>
		<li>Using the electrode&#39;s tip, carefully hook the EUS muscle bilaterally. Avoid punching too deep, as this may harm the muscle, which could lead to possible urine leakage.</li>
		<li>Use the 3-0 nonabsorbable monofilament to suture the pelvic and abdomen muscles and skin.</li>
	</ol>
	</li>
	<li>Waterproofing the skin
	<ol>
		<li>Apply a thin layer of cyanoacrylate glue to the skin where the electrodes exit to fix the electrodes in place.</li>
		<li>Apply the cyanoacrylate glue 1 cm away surrounding the urethral meatus and 3 cm further extending to the abdomen and sutured region. To avoid contact with the glue, carefully hold the meatus up with forceps.</li>
		<li>Use a 0.5-10 &#181;L micropipette to withdraw the accelerator liquid to dry the glue. 
		CAUTION: Accelerator liquid is a combustible liquid.</li>
		<li>Add the accelerator liquid to ensure proper adhesive reaction. This will help to dry the glue more quickly and ensure that it sets securely.</li>
	</ol>
	</li>
	<li>Urodynamic preparation
	<ol>
		<li>Prepare an inverted polystyrene weighing boat 4.5 cm in length, width, and depth. Cut it into a triangle shape with a base of 4 cm to put the mouse&#39;s urethra meatus in this space. Put the disposable base mold, 37 mm x 24 mm x 5 mm, under the space for collecting the urine.</li>
		<li>Reposition the mouse in a prone position and carefully move it onto a custom-made plate equipped with a gas mask.</li>
		<li>Ensure that the urethral meatus is properly positioned within the groove. After gently restraining the head and limbs of the mouse with tape, ensuring that the chest and nose are unobstructed for breathing, remove the inhalant anesthesia. Use pulse oximetry or monitor chest wall movement to check respiration. Place the plate on a heating pad for heat support and allow the mouse to recover until it regains full consciousness (Figure 2).</li>
		<li>Perform cystometry only when the mouse is fully awake, which is at least 40 min after recovery from anesthesia.</li>
	</ol>
	</li>
</ol>
7. Cystometry and EUS-EMG recording preparation
<ol>
	<li>Set up and calibrate the infusion pump according to the manufacturer&#39;s instructions.</li>
	<li>Take a 20 mL syringe with a diameter of 19.05 mm and fill it with sterile 0.9% NaCl at room temperature. Secure the syringe to the infusion pump. Set the infusion speed to 0.01 mL/min.</li>
	<li>Connect the syringe by the PE-30 tube to one side of the three-way connector. Connect the bladder catheter to the other side to a pressure transducer. Before connecting the bladder catheter, make sure to remove all air bubbles.</li>
	<li>Fix the pressure transducer at the same level as the mouse bladder. The pressure transducer is connected via an amplifier to the data acquisition system.</li>
	<li>Attach one ground line hook to the skin and the other to the electrode connector sites. Record the pressure in the software.</li>
	<li>After starting the software, check the intravesical pressure (IVP) and EUS-EMG signals. Save the sample name and set the time.</li>
	<li>Start the pump infusion. Record the signals for 40 minutes. At the end of the recording, euthanize the animal, as this is an end-stage procedure.</li>
</ol>

Precise Urodynamic Measurement in Awake Mice Using Cyanoacrylate Skin Barriers

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O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Best+practices+for+cystometric+evaluation+of+lower+urinary+tract+function+in+muriform+rodents.">Best practices for cystometric evaluation of lower urinary tract function in muriform rodents.</a> Neurourol Urodyn. 39 (6), 1868-1884 (2020).</li><li>Hashimoto, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sex+differences+in+lower+urinary+tract+function+in+mice+with+or+without+spinal+cord+injury.">Sex differences in lower urinary tract function in mice with or without spinal cord injury.</a> Neurourol Urodyn. 43 (1), 267-275 (2024).</li><li>Kadekawa, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+bladder+and+external+urethral+activity+in+mice+with+or+without+spinal+cord+injury-a+comparison+study+with+rats.">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>Lee, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effects+of+periurethral+muscle-derived+stem+cell+injection+on+leak+point+pressure+in+a+rat+model+of+stress+urinary+incontinence.">The effects of periurethral muscle-derived stem cell injection on leak point pressure in a rat model of stress urinary incontinence.</a> Int Urogynecol J. 14, 31-37 (2003).</li><li>Mann-Gow, T. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluating+the+procedure+for+performing+awake+cystometry+in+a+mouse+model.">Evaluating the procedure for performing awake cystometry in a mouse model.</a> J Vis Exp. (123), e55588 (2017).</li><li>Saito, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Time-dependent+progression+of+neurogenic+lower+urinary+tract+dysfunction+after+pinal+cord+injury+in+the+mouse+model.">Time-dependent progression of neurogenic lower urinary tract dysfunction after pinal cord injury in the mouse model.</a> Am J Physiol Renal Physioly. 321 (1), F26-F32 (2021).</li><li>Schneider, M. P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+urodynamic+model+for+lower+urinary+tract+assessment+in+awake+rats.">A novel urodynamic model for lower urinary tract assessment in awake rats.</a> BJU Int. 115, 8-15 (2015).</li><li>Habib, A., Mehanna, A., Medra, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cyanoacrylate:+A+handy+tissue+glue+in+maxillofacial+surgery:+Our+experience+in+alexandria,+egypt.">Cyanoacrylate: A handy tissue glue in maxillofacial surgery: Our experience in alexandria, egypt.</a> J Maxillofac Oral Surg. 12, 243-247 (2013).</li><li>Sunjic Roguljic, V., Roguljic, L., Jukic, I., Kovacic, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+influence+of+wound+closure+techniques+after+surgical+decompression+in+patients+with+carpal+tunnel+syndrome+on+sleep+disturbance+and+life+quality:+A+prospective+comparison+of+surgical+techniques.">The influence of wound closure techniques after surgical decompression in patients with carpal tunnel syndrome on sleep disturbance and life quality: A prospective comparison of surgical techniques.</a> Clin Pract. 14 (2), 546-555 (2024).</li><li>Sohn, J. 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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Muro-neuro-urodynamics;+a+review+of+the+functional+assessment+of+mouse+lower+urinary+tract+function.">Muro-neuro-urodynamics; a review of the functional assessment of mouse lower urinary tract function.</a> Front Physiol. 8, 240395 (2017).</li><li>Abdelkhalek, A. S., Youssef, H. A., Saleh, A. S., Bollen, P., Zvara, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anesthetic+protocols+for+urodynamic+studies+of+the+lower+urinary+tract+in+small+rodents-a+systematic+review.">Anesthetic protocols for urodynamic studies of the lower urinary tract in small rodents-a systematic review.</a> PloS One. 16 (6), e0253192 (2021).</li><li>Saab, B. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Short-term+memory+impairment+after+isoflurane+in+mice+is+prevented+by+the+&#945;5+&#947;-aminobutyric+acid+type+a+receptor+inverse+agonist+l-655,708.">Short-term memory impairment after isoflurane in mice is prevented by the &#945;5 &#947;-aminobutyric acid type a receptor inverse agonist l-655,708.</a> J Am Soc Anesthesiol. 113 (5), 1061-1071 (2010).</li><li>Cannon, T. W., Damaser, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+anesthesia+on+cystometry+and+leak+point+pressure+of+the+female+rat.">Effects of anesthesia on cystometry and leak point pressure of the female rat.</a> Life Sci. 69 (10), 1193-1202 (2001).</li><li>Weiss, D. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphology+of+the+external+genitalia+of+the+adult+male+and+female+mice+as+an+endpoint+of+sex+differentiation.">Morphology of the external genitalia of the adult male and female mice as an endpoint of sex differentiation.</a> Mol Cell Endocrinol. 354 (1-2), 94-102 (2012).</li><li>Leggat, P. A., Kedjarune, U., Smith, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Toxicity+of+cyanoacrylate+adhesives+and+their+occupational+impacts+for+dental+staff.">Toxicity of cyanoacrylate adhesives and their occupational impacts for dental staff.</a> Ind Health. 42 (2), 207-211 (2004).</li></ol>

The authors have nothing to disclose.

This urodynamic technique describes an improved procedure for measuring urine volume and EUS- EMG signal in awake and restrained mice. The presence of fur around the urethral meatus and abdominal area can interfere with the accuracy of the VV measurement by absorbing urine. Although the fur surrounding the urethral meatus and abdomen had been carefully shaved before surgery, the remaining small furs within these areas and the skin still absorbed urine, usually leaving a wet area in the abdomen after recording. This issue...

The storage and release of urine are dependent on the coordinated activity of the urinary bladder and external urethral sphincter (EUS). In some pathologies such as neurogenic bladder, both the bladder detrusor muscles and the sphincter can become dysfunctional, leading to significant bladder problems, especially after traumatic spinal cord injury(SCI)1.
Small rodents are commonly used as an experimental model to study the preclinical function of the lower urinary tract (LUT)2. Filling Cystometry (FC) and EUS electromyography (EUS-EMG) recording techniques can provide precise objective information depending on the choice of methods, accurate measurement, and interpretation of results3. Urodynamic tests are commonly used to evaluate the voiding volume (VV), voiding efficiency (VE), and bladder capacity4. VE measures how effectively the bladder can empty itself. It is calculated by dividing the voided volume by the sum of voided and residual volumes (VV+RV). On the other hand, bladder capacity is calculated by adding the VV (the amount of urine expelled during urination) to the RV (the amount of urine left in the bladder after urination)5. Therefore, the measurement of VV and RV are the keys to deducing other parameters.
Precisely measuring VV in mice during urodynamic tests presents various challenges. The urine of rodents, when physically restrained in a prone position, tends to be drawn downwards through the ventral abdominal wall due to the influence of gravity6. This phenomenon can lead to the absorption of urine by the abdominal fur and skin, which, in turn, underestimates the volume of urine excreted. Considering the small amount of urine produced by mouse, the impact of this absorbance on the accuracy of results is even more pronounced7. Furthermore, in models of SCI, VV is often lower than in normal mice due to the impact of the detrusor sphincter dyssynergia (DSD), which increases the risk of leak point pressures and urine absorption by the fur8. These factors have a significant impact on the results. Therefore, accurate measurement of VV and RV during terminal urodynamic studies in mice is crucial9. Currently, there is a lack of details in the methodologies provided in published literature about how to measure urine volume accurately in mouse models.
Cyanoacrylate adhesive is a type of glue that is commonly used in surgical procedures in human and animal models due to its quick and effective bonding properties10,11,12. This adhesive is particularly useful for closing wounds and lacerations, as it forms a strong and flexible bond when applied to the skin13. Moreover, it can be a great barrier against urine and wetness that may come into contact with fur and wounds11.
In this article, we have developed a novel and cost-effective technique that utilizes cyanoacrylate adhesive to achieve precise results in cystometry and EUS-EMG recordings in awake mice. This method will be beneficial in understanding the underlying causes of bladder dysfunction and devising more effective treatments for LUT disorders.

<table><tbody><tr><td>Accelerator</td><td>BOB SMITH INDUSTRIES</td><td>BSI-152</td><td></td></tr><tr><td>Cyanoacrylate&amp;nbsp;</td><td>TED PELLA, Inc</td><td>14478</td><td></td></tr><tr><td>Disposable base mold</td><td>TED PELLA, Inc</td><td>27147-4</td><td></td></tr><tr><td>Infusion pump</td><td>Harvard Apparatus PHD ULTRA</td><td>70-3006</td><td></td></tr><tr><td>Isoflurane</td><td>Henry Schein Inc</td><td>1182097</td><td></td></tr><tr><td>PIN</td><td>World Precision Instruments</td><td>5482</td><td></td></tr><tr><td>Polyethylene Tubing 30</td><td>Braintree Scientific Inc</td><td>PE30</td><td></td></tr><tr><td>Sterile Weighing Boat</td><td>HEATHROW SCIENTIFIC</td><td>797CK2</td><td></td></tr><tr><td>Windaq/Lite&amp;nbsp;</td><td>DATAQ INSTRUMENTS</td><td>249022</td><td></td></tr></tbody></table>

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.

Cystometry and EUS-EMG activity tracings were used to analyze the data. The continuous cystometry method involves the infusion of saline into the bladder and simultaneously measuring the pressure and volume changes in the bladder. To measure VV, 0.4 mL of saline was infused at a speed of 0.01 mL/min, and urine was collected over 40 min in a cap. The post-void residual (PVR) can be obtained by aspirating the saline through the catheter. In normal mice without glue, the sum of VV and RV was often less than 0.4 mL. After th...

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.

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 ...

optimizing-mouse-urodynamic-techniques-for-improved-accuracy

Research

JoVE Journal

Neuroscience

601 Views.  Indiana University School of Medicine. Het doel van deze studie is om de urodynamische technologie te verbeteren om de nauwkeurigheid van de urinemeting te vergroten. Nauwkeurige meting van de urineparameter wakkere muizen is cruciaal om disfunctie van de lagere urinewegen te begrijpen, met name bij aandoeningen zoals een neurogene blaas, posttraumatisch ruggenmergletsel. We hebben een unieke methode ontwikkeld met behulp van cyanoacrylaatlijm om een waterdichte huidbarrière rond de buik te creë...

Video: Auteur in de schijnwerpers: verbeterde urodynamische methode voor nauwkeurige urinemeting bij wakkere muizen met neurogene blaas

Transurethral Induction of Mouse Urinary Tract Infection

Uropathogenic bacterial strains of interest are grown on agar. Generally, uropathogenic E. coli (UPEC) and other strains can be grown overnight on Luria-Bertani (LB) agar at 37&deg;C in ambient air. UPEC strains grow as yellowish-white translucent colonies on LB agar. Following confirmation of appropriate colony morphology, single colonies are then picked to be cultured in broth. LB broth can be used for most uropathogenic bacterial strains. Two serial, overnight LB broth cultures can be employed to enhance expression of type I pili, a well-defined virulence factor for uropathogenic bacteria. Broth cultures are diluted to the desired concentration in phosphate buffered saline (PBS). Eight to 12 week old female mice are placed under isoflurane anesthesia and transurethrally inoculated with bacteria using polyethylene tubing-covered 30 gauge syringes. Typical inocula, which must be empirically determined for each bacterial/mouse strain combination, are 106 to 108 cfu per mouse in 10 to 50 microliters of PBS. After the desired infection period (one day to several weeks), urine samples and the bladder and both kidneys are harvested. Each organ is minced, placed in PBS, and homogenized in a Blue Bullet homogenizer. Urine and tissue homogenates are serially diluted in PBS and cultured on appropriate agar. The following day, colony forming units are counted.

This video will demonstrate methods to transurethrally induce mouse urinary tract infections and quantify the extent of resulting infections.

Uropathogenic bacterial strains of interest are grown on agar. Generally, uropathogenic E. coli (UPEC) and other strains can be grown overnight on ...

Immunology and Infection

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 ...

Medicine

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.

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 ...

Ultrasonography of the Adult Male Urinary Tract for Urinary Functional Testing

The incidence of clinical benign prostatic hyperplasia (BPH) and lower urinary tract symptoms (LUTS) is increasing due to the aging population, resulting in a significant economic and quality of life burden. Transgenic and other mouse models have been developed to recreate various aspects of this multifactorial disease; however, methods to accurately quantitate urinary dysfunction and the effectiveness of new therapeutic options are lacking. Here, we describe a method that can be used to measure bladder volume and detrusor wall thickness, urinary velocity, void volume and void duration, and urethral diameter. This would allow for the evaluation of disease progression and treatment efficacy over time. Mice were anesthetized with isoflurane, and the bladder was visualized by ultrasound. For non-contrast imaging, a 3D image was taken of the bladder to calculate volume and evaluate shape; the bladder wall thickness was measured from this image. For contrast-enhanced imaging, a catheter was placed through the dome of the bladder using a 27-gauge needle connected to a syringe by PE50 tubing. A bolus of 0.5 mL of contrast was infused into the bladder until a urination event occurred. Urethral diameter was determined at the point of the Doppler velocity sample window during the first voiding event. Velocity was measured for each subsequent event yielding a flow rate. In conclusion, high frequency ultrasound proved to be an effective method for assessing bladder and urethral measurements during urinary function in mice. This technique may be useful in the assessment of novel therapies for BPH/LUTS in an experimental setting.

We describe the use of high frequency ultrasound with contrast imaging as a method to measure bladder volume, bladder wall thickness, urine velocity, void volume, void duration, and urethral diameter. This strategy can be used to assess voiding dysfunction and treatment efficacy in various mouse models of lower urinary tract dysfunction (LUTD).

The incidence of clinical benign prostatic hyperplasia (BPH) and lower urinary tract symptoms (LUTS) is increasing due to the aging population, resulting ...

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. ...

Patient-derived Tumor Xenografting: A Technique to Generate Experimental Mouse Model for Evaluating Urothelial Cell Carcinoma

Source: Moret, R. et al. <a href="https://www.jove.com/video/59223" target="_blank">Patient-derived Orthotopic Xenograft Models for Human Urothelial Cell Carcinoma and Colorectal Cancer Tumor Growth and Spontaneous Metastasis.</a> J. Vis. Exp. (2019)
This video describes the procedure to generate an experimental xenograft murine model, which can be used for studying urothelial cell carcinoma progression.

Source: Moret, R. et al. Patient-derived Orthotopic Xenograft Models for Human Urothelial Cell Carcinoma and Colorectal Cancer Tumor Growth and ...

JoVE Encyclopedia of Experiments

Cancer Research

Urinary Tract Cancer

In vivo studies

Unilateral Ureteral Obstruction Murine Model: A Surgical Procedure to Obstruct the Urinary Flow from a Kidney in Mouse Models

Source: Yanai, K. et al. <a href="https://www.jove.com/video/61383" target="_blank">Quantitative real-time PCR evaluation of microRNA expressions in mouse kidney with unilateral ureteral obstruction.</a> J. Vis. Exp. (2020)
This video demonstrates the procedure to generate a unilateral ureteral obstruction mouse model. The procedure involves cutting one of the ureters of the mouse to obstruct urine flow.

Source: Yanai, K. et al. Quantitative real-time PCR evaluation of microRNA expressions in mouse kidney with unilateral ureteral obstruction. J. Vis. Exp. ...

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/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.

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

Source: Codolo, G. et al. <a href="https://app.jove.com/v/52743">Evaluation of the Efficacy of the H. pylori Protein HP-NAP as a Therapeutic Tool for Treatment of Bladder Cancer in an Orthotopic Murine Model.</a>&#160;J. Vis. Exp. (2015)
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 na&#239;ve 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 destruction, reducing tumor volume.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Optimizing Mouse Urodynamic Techniques for Improved Accuracy

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