Optimizing Mouse Urodynamic Techniques for Improved Accuracy

Summary

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.

Abstract

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.

Introduction

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 properties^10,11,12. 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.

Protocol

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

1. 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.
2. 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.
3. 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.

2. Preparation of electrodes

1. 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.
2. 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.
3. 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.

3. Preparation of animal

1. 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

4. Anesthesia during surgery

1. Place the animals in a 2% isoflurane and pure oxygen (1 L/min) chamber. Confirm the animal's complete anesthesia using negative toe-pinch examination before transferring it to the mask. Once confirmed, change the gas condition to a mask.
2. 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.

5. Surgical preparation

1. 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). Administer 3 mg/kg Meloxicam (5 mg/ml) subcutaneously as a pre-emptive analgesic to the mouse before surgery to ensure effective pain management.
2. 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.
3. Autoclave all surgical tools and electrode wires at 121°C for 15–20 minutes to ensure sterility prior to the procedure. Soak the polyethylene tube in a glutaraldehyde or ethylene oxide for 20–30 minutes, then rinse thoroughly with sterile saline to remove any residues

6. Surgical procedure

1. Bladder catheter implantation
   1. 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.
   2. 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.
   3. Grasp the bladder dome and place a purse string by utilizing a 5-0 nonabsorbable monofilament suture with a taper-tip needle.
   4. Using micro scissors, create a small cystostomy within the purse string and make a hole until urine flows out.
   5. 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.
   6. 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.
   7. Once the saline comes out of the urethra, withdraw the saline to decompress the bladder.
2. EUS electrodes implantation (Figure 1)
   1. Use surgical scissors to extend the abdominal incision up to the pelvic floor.
   2. 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.
   3. Puncture the skin bilaterally, 1 cm away from the urethral meatus, using the needle containing the electrode.
   4. Carefully grab the hook tip with forceps and gently pull the needle away from the skin.
   5. Using the electrode'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.
   6. Use the 3-0 nonabsorbable monofilament to suture the pelvic and abdomen muscles and skin.
3. Waterproofing the skin
   1. Apply a thin layer of cyanoacrylate glue to the skin where the electrodes exit to fix the electrodes in place.
   2. 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.
   3. Use a 0.5-10 µL micropipette to withdraw the accelerator liquid to dry the glue.
      CAUTION: Accelerator liquid is a combustible liquid.
   4. Add the accelerator liquid to ensure proper adhesive reaction. This will help to dry the glue more quickly and ensure that it sets securely.
4. Urodynamic preparation
   1. 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'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.
   2. Reposition the mouse in a prone position and carefully move it onto a custom-made plate equipped with a gas mask.
   3. 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).
   4. Perform cystometry only when the mouse is fully awake, which is at least 40 min after recovery from anesthesia.

7. Cystometry and EUS-EMG recording preparation

1. Set up and calibrate the infusion pump according to the manufacturer's instructions.
2. 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.
3. 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.
4. 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.
5. Attach one ground line hook to the skin and the other to the electrode connector sites. Record the pressure in the software.
6. After starting the software, check the intravesical pressure (IVP) and EUS-EMG signals. Save the sample name and set the time.
7. 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.

Results

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'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 signals^5,14, 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.

Figure 1: Displacement of the electromyography electrodes. Implantation of electrodes (yellow asterisk) bilaterally to the external urethral muscle (EUS; black arrows). Please click here to view a larger version of this figure.

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. Please click here to view a larger version of this figure.

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. Please click here to view a larger version of this figure.

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). Please click here to view a larger version of this figure.

Discussion

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.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Accelerator	BOB SMITH INDUSTRIES	BSI-152
Cyanoacrylate	TED PELLA, Inc	14478
Disposable base mold	TED PELLA, Inc	27147-4
Infusion pump	Harvard Apparatus PHD ULTRA	70-3006
Isoflurane	Henry Schein Inc	1182097
PIN	World Precision Instruments	5482
Polyethylene Tubing 30	Braintree Scientific Inc	PE30
Sterile Weighing Boat	HEATHROW SCIENTIFIC	797CK2
Windaq/Lite	DATAQ INSTRUMENTS	249022