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. J., Gruber, T. M., Zahorsky-Reeves, J. L., Lawson, G. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+2-ethyl-cyanoacrylate+and+2-butyl-cyanoacrylate+for+use+on+the+calvaria+of+cd1+mice.">Comparison of 2-ethyl-cyanoacrylate and 2-butyl-cyanoacrylate for use on the calvaria of cd1 mice.</a> J Am Assoc Lab Anim Sci. 55 (2), 199-203 (2016).</li><li>Ren, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Injectable,+self-healing+hydrogel+adhesives+with+firm+tissue+adhesion+and+on-demand+biodegradation+for+sutureless+wound+closure.">Injectable, self-healing hydrogel adhesives with firm tissue adhesion and on-demand biodegradation for sutureless wound closure.</a> Sci Adv. 9 (33), 4327 (2023).</li><li>Ito, H., Pickering, A. E., Kanai, A., Fry, C. H., Drake, M. 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 is particularly noticeable in female rodents due to the extremely short distance between the urethral meatus and the surrounding skin18. In this technique, cyanoacrylate glue was applied on the abdominal and surrounding urethral skin to create a waterproof skin surface and provide a precise assessment of urinary volume during the urodynamic recording, allowing for a better understanding of bladder function. The glue was applied with precision, ensuring that it covered the skin surrounding the meatus and nearby. The purpose of applying the glue was to create a waterproof barrier that would prevent the furs from absorbing any urine. The glue was spread evenly, with care taken to avoid any clumping or blocking of the urethral meatus. The recorded results of the procedure confirmed that our objective had been completely achieved, as the sum of VV and RV remained constant at infusion volume, and no further wet areas were observed. To ensure the accuracy of measurements, we checked the bladder after the experiment, and it was found to be empty. This additional step of checking the bladder is crucial as it eliminates any possibility of urine retention, causing a discrepancy between the amount we withdrew through a syringe and the actual amount of RV.
This method has limitations: 1) it is unsuitable for longitudinal and multiple-time-point studies. 2) it cannot be applied to a freely moving mouse. 3) if the detachment of the electrodes from EUS occurs, it is difficult to open the abdomen and reinstall them. 4) While cyanoacrylate adhesives are a valuable tool in many surgical settings due to their ease of use and effectiveness, it is important to use them cautiously and follow proper safety protocols to minimize any potential risks. Cyanoacrylate is generally safe for the skin, but frequent contact with it should be avoided, and researchers should take appropriate personal protective measures. Cyanoacrylate adhesives can release toxic vapors if inhaled. To minimize the risk of inhaling these vapors, researchers should maintain higher levels of humidity and optimize room ventilation in the working environment19. Special air conditioning filters can also be used to further reduce the toxicity of the vapors.
Overall, this experiment provided important insights into the accuracy of measuring voided urine during the urodynamic recording and helped to identify potential sources of error that could have led to discrepancies in the total amount of VV and RV after infusion.

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Accelerator</td>
			<td>BOB SMITH INDUSTRIES</td>
			<td>BSI-152</td>
			<td></td>
		</tr>
		<tr>
			<td>Cyanoacrylate&#160;</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&#160;</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 the experiment, the fur in the abdomen and surrounding the meatus was wet due to the absorption of urine (Figure 3A). After applying a thin layer of glue to cover small furs, the sum of VV and RV was shown to be 0.4 mL, and there was no wet area (Figure 3B,C).
The resulting cystometry tracings provided a detailed analysis of various parameters, including maximum voiding bladder contraction pressure (27.2 cmH2O), contraction duration (16.26 s), and inter-contraction interval (4.48 min). At the same time, we had a good recording of intravesical pressure and EUS-EMG signals in mice, as shown in Figure 4.
Many mouse urodynamic measurements are performed under anesthesia14. Although this may seem like a convenient method to reduce the noise of electrical signals and loss of urine resulting from the animal&#39;s movement, it is essential to consider that the anesthetic drugs can affect the urinary flow, which may lead to inaccurate or unreliable results15. Therefore, urodynamic recording in awake animals is more popular to obtain results closer to the physiological condition. The urodynamic recording in awake animals usually begins after a 40-50 min period of recovery from isoflurane16. This process involves closely monitoring the mice to ensure that they are relaxed and comfortable without the need for anesthesia. It has been observed through several experiments that the movement of a conscious mouse can affect urodynamic signals5,14, leading to inaccurate measurements of specific parameters such as leak point pressure, VV, and VE17. As a result, we have implemented a method by partially restraining conscious mice to ensure more reliable urodynamic results. However, even with limited restraint, the conscious mice still struggle when they wake up immediately from the anesthesia, which can also cause detachment or unstable contact between the electrode hook and the EUS and create a significant noise in the EUS-EMG signals. As shown in Figure 3B, to minimize these artifacts, we have taken the approach of fixing the electrodes with glue at the exit point from the skin. This method has proven to be effective in minimizing the movement of electrodes and the subsequent artifacts that they can produce.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67019/67019fig01.jpg" /> 
Figure 1: Displacement of the electromyography electrodes. Implantation of electrodes (yellow asterisk) bilaterally to the external urethral muscle (EUS; black arrows). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67019/67019fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67019/67019fig02.jpg" /> 
Figure 2: Restraint of the awake mouse. After implantation of catheter and electrodes, the mouse was restrained on the plate for stability during urodynamic recording. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67019/67019fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/67019/67019fig03.jpg" /> 
Figure 3: Abdominal and meatus regions after urodynamic recording. (A) A large wet area (contoured by a red dash line) was seen in the abdomen and genital regions. (B) Dry, waterproof abdominal and genital areas were created with cyanoacrylate glue (contoured by a red dash line) after recording. (C) A urine drop (yellow arrow) formed at the meatus during urodynamic recording and stayed as a drop for a long time without being absorbed by the skin and fur. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67019/67019fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/67019/67019fig04.jpg" /> 
Figure 4: Representative traces of cystometry and external urethral sphincter electromyography (EUS-EMG) in an awake and restrained female mouse. (A) Trace A: Simultaneous recordings of continuous cystometrogram (CMG) and EUS-EMG (upper and lower traces, respectively). (B) Trace B is the expanded portion of trace A, indicated by a rectangular box with different time scales. During the voiding phase, intermittent voiding coincided with reductions in intravesical pressure in the CMG recording (top trace; arrows), which occurred during low tonic and reduction periods of EUS-EMG activity (bottom trace; arrows). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67019/67019fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

Research

JoVE Journal

Neuroscience

540 Views.  Indiana University School of Medicine. 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.

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

Operant Sensation Seeking in the Mouse

Operant methods are powerful behavioral tools for the study of motivated behavior. These 'self-administration' methods have been used extensively in drug addiction research due to their high construct validity. Operant studies provide researchers a tool for preclinical investigation of several aspects of the addiction process. For example, mechanisms of acute reinforcement (both drug and non-drug) can be tested using pharmacological or genetic tools to determine the ability of a molecular target to influence self-administration behavior1-6. Additionally, drug or food seeking behaviors can be studied in the absence of the primary reinforcer, and the ability of pharmacological compounds to disrupt this process is a preclinical model for discovery of molecular targets and compounds that may be useful for the treatment of addiction3,7-9. One problem with performing intravenous drug self-administration studies in the mouse is the technical difficulty of maintaining catheter patency. Attrition rates in these experiments are high and can reach 40% or higher10-15. Another general problem with drug self-administration is discerning which pharmacologically-induced effects of the reinforcer produce specific behaviors. For example, measurement of the reinforcing and neurological effects of psychostimulants can be confounded by their psychomotor effects. Operant methods using food reinforcement can avoid these pitfalls, although their utility in studying drug addiction is limited by the fact that some manipulations that alter drug self-administration have a minimal impact on food self-administration. For example, mesolimbic dopamine lesion or knockout of the D1 dopamine receptor reduce cocaine self-administration without having a significant impact on food self-administration 12,16. 
Sensory stimuli have been described for their ability to support operant responding as primary reinforcers (i.e. not conditioned reinforcers)17-22. Auditory and visual stimuli are self-administered by several species18,21,23, although surprisingly little is known about the neural mechanisms underlying this reinforcement. The operant sensation seeking (OSS) model is a robust model for obtaining sensory self-administration in the mouse, allowing the study of neural mechanisms important in sensory reinforcement24. An additional advantage of OSS is the ability to screen mutant mice for differences in operant behavior that may be relevant to addiction. We have reported that dopamine D1 receptor knockout mice, previously shown to be deficient in psychostimulant self-administration, also fail to acquire OSS24. This is a unique finding in that these mice are capable of learning an operant task when food is used as a reinforcer. While operant studies using food reinforcement can be useful in the study of general motivated behavior and the mechanisms underlying food reinforcement, as mentioned above, these studies are limited in their application to studying molecular mechanisms of drug addiction. Thus, there may be similar neural substrates mediating sensory and psychostimulant reinforcement that are distinct from food reinforcement, which would make OSS a particularly attractive model for the study of drug addiction processes. The degree of overlap between other molecular targets of OSS and drug reinforcers is unclear, but is a topic that we are currently pursuing. While some aspects of addiction such as resistance to extinction may be observed with OSS, we have found that escalation 25 is not observed in this model24. Interestingly, escalation of intake and some other aspects of addiction are observed with self-administration of sucrose26. Thus, when non-drug operant procedures are desired to study addiction-related processes, food or sensory reinforcers can be chosen to best fit the particular question being asked. 
In conclusion, both food self-administration and OSS in the mouse have the advantage of not requiring an intravenous catheter, which allows a higher throughput means to study the effects of pharmacological or genetic manipulation of neural targets involved in motivation. While operant testing using food as a reinforcer is particularly useful in the study of the regulation of food intake, OSS is particularly apt for studying reinforcement mechanisms of sensory stimuli and may have broad applicability to novelty seeking and addiction.

In this protocol we describe a method of operant learning using sensory stimuli as a reinforcer in the mouse. It requires no prior training or food restriction, and it allows the study of motivated behavior without the use of a pharmacological or natural reinforcer such as food.

Operant methods are powerful behavioral tools for the study of motivated behavior.  These 'self-administration' methods have been used extensively in drug ...

Multiple-mouse Neuroanatomical Magnetic Resonance Imaging

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and evaluating mouse models of human disease. MRI is an excellent modality for investigating genetically altered animals. It is capable of whole brain coverage, can be used in vivo, and provides multiple contrast mechanisms for investigating different aspects of neuranatomy and physiology. The advent of high-field scanners along with the ability to scan multiple mice simultaneously allows for rapid phenotyping of novel mutations.
Effective mouse MRI studies require attention to many aspects of experiment design. In this article, we will describe general methods to acquire quality images for mouse phenotyping using a system that images mice concurrently in shielded transmit/receive radio frequency (RF) coils in a common magnet (Bock et al., 2003). We focus particularly on anatomical phenotyping, an important and accessible application that has shown a high potential for impact in many mouse models at our imaging centre. Before we can provide the detailed steps to acquire such images, there are important practical considerations for both in vivo brain imaging (Dazai et al., 2004) and ex vivo brain imaging (Spring et al., 2007) that should be noted. These are discussed below.

Magnetic resonance imaging (MRI) has become an increasingly popular tool for examining the phenotype of genetically altered mice. This article illustrates the methods necessary to achieve high-throughput phenotyping of genetically altered mice using multiple-mouse MRI.

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and ...

Improved Preparation and Preservation of Hippocampal Mouse Slices for a Very Stable and Reproducible Recording of Long-term Potentiation

Long-term potentiation (LTP) is a type of synaptic plasticity characterized by an increase in synaptic strength and believed to be involved in memory encoding. LTP elicited in the CA1 region of acute hippocampal slices has been extensively studied. However the molecular mechanisms underlying the maintenance phase of this phenomenon are still poorly understood. This could be partly due to the various experimental conditions used by different laboratories. Indeed, the maintenance phase of LTP is strongly dependent on external parameters like oxygenation, temperature and humidity. It is also dependent on internal parameters like orientation of the slicing plane and slice viability after dissection.
The optimization of all these parameters enables the induction of a very reproducible and very stable long-term potentiation. This methodology offers the possibility to further explore the molecular mechanisms involved in the stable increase in synaptic strength in hippocampal slices. It also highlights the importance of experimental conditions in in vitro investigation of neurophysiological phenomena.

This paper presents a complete methodology to prepare and preserve in vitro acute hippocampal slices from adult mice. This protocol allows the recording of very stable long-lasting long-term potentiation (LTP) for more than 8 hours with a success rate of 95%.

Long-term potentiation (LTP) is a type of synaptic plasticity characterized by an increase in synaptic strength and believed to be involved in memory ...

Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques

The retina is the gateway to the visual system. To understand visual signal processing mechanisms, we investigate retinal neural network functions. Retinal neurons in the network comprise of numerous subtypes. More than 10 subtypes of bipolar cells, ganglion cells, and amacrine cells have been identified by morphological studies. Multiple subtypes of retinal neurons are thought to encode distinct features of visual signaling, such as motion and color, and form multiple neural pathways. However, the functional roles of each neuron in visual signal processing are not fully understood. The patch clamp method is useful to address this fundamental question. Here, a protocol to record light-evoked synaptic responses in mouse retinal neurons using patch clamp recordings in dark-adapted conditions is provided. The mouse eyes are dark-adapted O/N, and retinal slice preparations are dissected in a dark room using infrared illumination and viewers. Infrared light does not activate mouse photoreceptors and thus preserves their light responsiveness. Patch clamp is used to record light-evoked responses in retinal neurons. A fluorescent dye is injected during recordings to characterize neuronal morphological subtypes. This procedure enables us to determine the physiological functions of each neuron in the mouse retina.

We will demonstrate how to prepare retinal slices from the mouse eye and record light responses in retinal neurons. The entire procedure is conducted in dark-adapted conditions.

The retina is the gateway to the visual system. To understand visual signal processing mechanisms, we investigate retinal neural network functions. ...

Piezo High Accuracy Surgical Osteal Removal (PHASOR): A Technique for Improved Cranial Window Surgery in Mice

Multiphoton microscopy has been widely adapted for imaging neurons in vivo. Repeated imaging requires implantation of a cranial window or repeated thinning of the skull. Cranial window surgery is typically performed with a high speed rotary drill, and many investigators find it challenging to prevent the drill from damaging the delicate dura and blood vessels. Extensive training and practice is required to remove the bone without damage to underlying tissue and thus cranial window surgery can be difficult, time consuming, and produce tissue damage. Piezoelectric surgery, which is extensively used for maxillofacial and dental surgery, utilizes ultrasonic vibrations to remove bone without damaging soft tissues. We have developed a method applying piezoelectric surgery to improve cranial window surgery in mice in preparation for multiphoton imaging. Comparisons within our lab find that the method requires less surgery time and has a lower average rate of complications due to dural bleeding than cranial window surgery with a rotary drill.

Piezo High Accuracy Surgical Osteal Removal PHASOR: A Technique for Improved Cranial Window Surgery in Mice

Piezoelectric surgery has led to improvements in human maxillofacial and dental surgery. We have developed a protocol to optimize piezoelectric surgery for cranial window surgery in mice.

Multiphoton microscopy has been widely adapted for imaging neurons in vivo. Repeated imaging requires implantation of a cranial window or repeated ...

An Improved Method for Collection of Cerebrospinal Fluid from Anesthetized Mice

The cerebrospinal fluid (CSF) is a valuable body fluid for analysis in neuroscience research. It is one of the fluids in closest contact with the central nervous system and thus, can be used to analyze the diseased state of the brain or spinal cord without directly accessing these tissues. However, in mice it is difficult to obtain from the cisterna magna due to its closeness to blood vessels, which often contaminate samples. The area for CSF collection in mice is also difficult to dissect to and often only small samples are obtained (maximum of 5-7 &#181;L or less). This protocol describes in detail a technique that improves on current methods of collection to minimize contamination from blood and allow for the abundant collection of CSF (on average 10-15 &#181;L can be collected). This technique can be used with other dissection methods for tissue collection from mice, as it does not impact any tissues during CSF extraction. Thus, the brain and spinal cord are not affected with this technique and remain intact. With greater CSF sample collection and purity, more analyses can be used with this fluid to further aid neuroscience research and better understand diseases affecting the brain and spinal cord.

This protocol describes an improved technique for the abundant collection of cerebrospinal fluid (CSF) with no contamination from blood. With greater sample collection and purity, more analyses can be performed using CSF to further our understanding of diseases that affect the brain and spinal cord.

The cerebrospinal fluid (CSF) is a valuable body fluid for analysis in neuroscience research. It is one of the fluids in closest contact with the central ...

Construction of an Improved Multi-Tetrode Hyperdrive for Large-Scale Neural Recording in Behaving Rats

Monitoring the activity patterns of a large population of neurons over many days in awake animals is a valuable technique in the field of systems neuroscience. One key component of this technique consists of the precise placement of multiple electrodes into desired brain regions and the maintenance of their stability. Here, we describe a protocol for the construction of a 3D-printable hyperdrive, which includes eighteen independently adjustable tetrodes, and is specifically designed for in vivo extracellular neural recording in freely behaving rats. The tetrodes attached to the microdrives can either be individually advanced into multiple brain regions along the track, or can be used to place an array of electrodes into a smaller area. The multiple tetrodes allow for simultaneous examination of action potentials from dozens of individual neurons, as well as local field potentials from populations of neurons in the brain during active behavior. In addition, the design provides for simpler 3D drafting software that can easily be modified for differing experimental needs.

We present the construction of a 3D-printable hyperdrive with eighteen independently adjustable tetrodes. The hyperdrive is designed to record brain activity in freely behaving rats over a period of several weeks.

Monitoring the activity patterns of a large population of neurons over many days in awake animals is a valuable technique in the field of systems ...

Rabies Necropsy Techniques in Large and Small Animals

The New York State Department of Health (NYSDOH) Rabies Laboratory receives between 6,000 to 9,000 specimens annually and performs rabies testing for the entire state, with the exception of New York City. The Rabies laboratory necropsies a variety of animals ranging in size from bats to bovids. Most of these specimens are animals exhibiting neurological signs, however, less than 10% actually test positive for rabies; implying trauma, lesions or other infectious agents as the cause of these symptoms. Due to the risk of aerosolizing undiagnosed infectious agents, the Rabies Laboratory does not use power tools or saws. Three necropsy techniques will be presented for animals whose skulls are impenetrable with scissors. The laboratory has implemented these techniques to decrease potential exposure to infectious agents, eliminate unnecessary manipulation of the specimen and reduce processing time. The advantages of a preferred technique opposed to another is subject to the trained individual processing the specimen.

The goal of this protocol is to demonstrate safe necropsy techniques in small and large animals to obtain satisfactory tissue samples for rabies testing.

The New York State Department of Health (NYSDOH) Rabies Laboratory receives between 6,000 to 9,000 specimens annually and performs rabies testing for the ...

Generation of Oligodendrocytes and Oligodendrocyte-Conditioned Medium for Co-Culture Experiments

In the central nervous system, oligodendrocytes are well-known for their role in axon myelination, that accelerates the propagation of action potentials through saltatory conduction. Moreover, an increasing number of reports suggest that oligodendrocytes interact with neurons beyond myelination, notably through the secretion of soluble factors. Here, we present a detailed protocol allowing purification of oligodendroglial lineage cells from glial cell cultures also containing astrocytes and microglial cells. The method relies on overnight shaking at 37 &#176;C, which allows selective detachment of the overlying oligodendroglial cells and microglial cells, and the elimination of microglia by differential adhesion. We then describe the culture of oligodendrocytes and production of oligodendrocyte-conditioned medium (OCM). We also provide the kinetics of OCM treatment or oligodendrocytes addition to purified hippocampal neurons in co-culture experiments, studying oligodendrocyte-neuron interactions.

Herein, we display an efficient method for the purification of oligodendrocytes and production of oligodendrocyte-conditioned medium that can be used for co-culture experiments.

In the central nervous system, oligodendrocytes are well-known for their role in axon myelination, that accelerates the propagation of action potentials ...

Adaptable Angled Stereotactic Approach for Versatile Neuroscience Techniques

Stereotactic surgery is an essential tool in the modern neuroscience lab. However, the ability to precisely and accurately target difficult-to-reach brain regions still presents a challenge, particularly when targeting brain structures along the midline. These challenges include avoiding of the superior sagittal sinus and third ventricle and the ability to consistently target selective and discrete brain nuclei. In addition, more advanced neuroscience techniques (e.g., optogenetics, fiber photometry, and two-photon imaging) rely on targeted implantation of significant hardware to the brain, and spatial limitations are a common hindrance. Presented here is a modifiable protocol for stereotactic targeting of rodent brain structures using an angled coronal approach. It can be adapted to 1) mouse or rat models, 2) various neuroscience techniques, and 3) multiple brain regions. As a representative example, it includes the calculation of stereotactic coordinates for targeting of the mouse hypothalamic ventromedial nucleus (VMN) for an optogenetic inhibition experiment. This procedure begins with the bilateral microinjection of an adeno-associated virus (AAV) encoding a light-sensitive chloride channel (SwiChR++) to a Cre-dependent mouse model, followed by the angled bilateral implantation of fiberoptic cannulae. Using this approach, findings show that activation of a subset of VMN neurons is required for intact glucose counterregulatory responses to insulin-induced hypoglycemia.

Described here is a stereotactic procedure that can target challenging and difficult-to-reach brain regions (due to spatial limitations) using an angled coronal approach. This protocol is adaptable to both mouse and rat models and can be applied to diverse neuroscientific applications, including cannula implantation and microinjections of viral constructs.

Stereotactic surgery is an essential tool in the modern neuroscience lab. However, the ability to precisely and accurately target difficult-to-reach brain ...