Evaluating the Procedure for Performing Awake Cystometry in a Mouse Model

Summary

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.

Abstract

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.

Introduction

Filling cystometry (FC) is a diagnostic method that involves placing a catheter into the urinary bladder to record pressure during slow bladder filling. First introduced in 1927 as a clinical diagnostic method to evaluate lower urinary tract function, it has remained widely used.¹ In research applications, FC can be used to test bladder function in healthy and diseased animal models and to study the effects of pharmacological agents. Rodent animal models are commonly used to investigate lower urinary tract function.² In this group of mammals, FC was first developed for use in rats.³ Here, the methodology to implant a tube into the urinary bladder and perform FC has been well described and used by many researchers with an acceptable level of reproducibility.⁴ The availability of transgenic and knock out strains make mice a valuable species for numerous research areas, including the field of lower urinary tract dysfunction. The methodology used for performing mouse cystometry varies appreciably between laboratories, making it difficult to compare results.⁵

Compared to ex vivo models, FC preserves lower urinary tract anatomy, allowing the coordinated function between the bladder and its outlet during the storage and voiding phases of the micturition cycle to be assessed. Previous research shows that numerous, commonly used anesthetics suppress micturition contraction. Agents that preserve urinary bladder smooth muscle contraction (urethane, α-chloralose, ketamine and xylazine), allowing the animal to micturate, still significantly reduce functional bladder capacity and suppress neurotransmission.^6,7,8,9 Although technically more challenging, FC performed in awake ambulating animals preserves the functional integrity of the micturition reflex.

Lower urinary tract function is influenced by multiple factors, including post-operative bladder wall swelling, stress due to pain and discomfort, and environmental influences. Using a surgical technique that minimizes tissue damage during tube implantation and recording methods that reduce tube movement, while simultaneously allowing the animal to ambulate freely, are essential for obtaining accurate and reproducible recordings.

If performed adequately, in vivo FC in freely moving animals can provide data that reliably reflects physiological bladder function.¹⁰ FC in freely moving animals can provide data on the following parameters; Basal or baseline pressure: Minimum pressure between two micturitions. Intermicturition pressure: Mean pressure between two micturitions. Threshold pressure: Intravesical pressure immediately before micturition. Maximum pressure: Maximum bladder pressure during a micturition cycle. Spontaneous activity (or mean intermicturition oscillatory pressure): Intermicturition pressure minus basal pressure. Non-voiding contractions: Increase in intravesical pressure during the filling phase, not associated with the release of fluid. Bladder compliance: Bladder capacity divided by threshold pressure minus basal pressure. Micturition frequency: Number of micturitions per unit of time. Intermicturition interval: Period between two maximum voiding pressures. Bladder capacity: Infused volume divided by the number of micturitions. A detailed description of these parameters and standardized terminology has been previously published.¹¹

FC can be performed using a continuous or single cycle intravesical infusion method. Continuous cystometry allows for recording of multiple micturition cycles and selecting representative data based on reproducibility. Its accuracy in measuring bladder capacity is limited due to the unknown residual volume. In addition, it is challenging to collect small voided volumes (which based on strain and sex vary between 30 and 184 µL) in freely ambulating mice. Using this method to record voided volume is less accurate compared to an anesthetized preparation, but it is superior in that it avoids the suppressive effects of anesthetics on bladder function. Single cycle cystometry should be used to assess bladder capacity. In this method, the bladder is emptied by aspiration prior to infusion and capacity is calculated as a function of infusion rate multiplied by time to maximum pressure.

Although the technique of performing cystometry in small rodents has been published, it described the surgery performed in a rat and recommended that mouse cystometry should be performed under urethane anesthesia.¹⁰ The goal of this communication is to describe both the microsurgical techniques used to implant an intravesical tube into the dome of the urinary bladder and the experimental set-up used to record lower urinary tract function, in vivo, during continuous bladder filling and micturition in an awake freely moving mouse. In addition, experiments were performed to address how tubing length, diameter, and material, as well as the methodology for performing in vivo FC, affect the recording. This experimental protocol summarizes previously published techniques and proposes a number of modifications based on experimental results.

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Protocol

Animals were housed in the University of Vermont Animal Care Facility according to institutional guidelines. All animal experiments were carried out in accordance with the National Institutes of Health guide for the care and use of laboratory animals.

1. Intravesical Tube Implantation

1. Preparation of tubing and instruments for the surgical procedure
   1. Cut a 7-cm piece of PE10 tubing to make the catheter for implantation.
   2. Create a flare at one end of the PE10 tube by slowly advancing the end towards an open flame.
      NOTE: Quickly withdraw the tube as soon as the flare develops.
   3. Apply three drops of all-purpose hot glue, using the low heat setting on a glue gun, at 4.5, 5, and 5.5 cm from the flared end on the outside of the PE10 tube. These will help secure the tube at the animal's back. (Figure 1)
   4. Sterilize the tubing by soaking it in 70% ethanol and then flush it with sterile 0.9% NaCl prior to use. Leave the tube filled to avoid introducing air bubbles into the system.
   5. Create a 30-gauge plug to seal the end of the PE10 catheter by separating a 30-gauge needle from the hub by manually manipulating the proximal end side-to-side. Apply a drop of hot glue to the end. Ensure that the seal is watertight. (Figure 2)
   6. Use the following microsurgical instruments: Two pairs of Dumont #7 curved microforceps, two pairs of Dumont #5 curved microforceps, a 21 G needle, ultrafine straight hemostat, micro scissors, small dissecting scissors, and a micro needle holder.
   7. Sterilize all instruments before starting the procedure.
2. Preparation of the animal
   1. After anesthetizing the animal, shave the lower half of the abdomen first, then turn the animal prone and shave and clean the area on the upper back with 70% alcohol followed by betadine. Apply vet ointment to the eyes to prevent dryness. Next, use a pair of straight, blunt scissors and pair of Dumont #7 curved microforceps to make a 1.5 cm long skin incision between the scapulae and place the animal supine on top of a heating pad (37 °C) covered with sterile drapes.
   2. Finally, clean the abdomen with alcohol and betadine.
3. Surgical procedure
   NOTE: Perform all surgical procedures under an operating microscope with magnification ranging from 3.15X to 20X. After placing the animal onto the sterile drapes, put on sterile gloves. Continue using sterile procedures throughout the entire surgery.
   1. Place the animal in an induction box and anesthetize using 2% inhaled isoflurane with an oxygen carrier (1 L/min). Maintain anesthesia throughout the procedure by placing the animal's head in a nose cone and using 2% inhaled isoflurane with an oxygen carrier (1 L/min). Begin the surgery after receiving a negative response from the toe-pinch test.
   2. Use a pair of straight, blunt scissors and pair of Dumont #7 curved microforceps to make a 1.5 cm lower, midline abdominal incision through the skin. Subsequently, create a matching incision through the fascia along the linea alba and muscle to expose the dome and upper half of the urinary bladder. Avoid injuring the bladder by applying upward traction to each tissue layer using a pair of Dumont #7 curved microforceps. Keep the abdominal viscera from desiccation by adding drops of warm physiological saline.
   3. Rotate the animal onto its side to access the incision on the nape of the neck. Push a narrow hemostat subcutaneously though the incision. The subcutaneous channel should start on the back, and continue along the side.
   4. Once the tip of the instrument reaches the bottom of the rib cage, turn the tip towards the midline and inside of the abdomen (there will be a slight pop when piercing the abdominal wall muscles). Continue advancing the hemostat until the tip is exposed at the abdominal incision under the muscular layer. (Figure 3)
   5. Grasp the "non-flared" end of the tubing with the hemostat and slowly retract the tool, pulling the end of the tube out via the incision at the back of the neck. Adjust the flared end of the tubing so that it lies directly above the dome of the bladder.
   6. Make a loose tie of 6-0 monofilament suture (non-absorbable) and place it on top of the bladder dome. This tie will be used later to secure the tube in the bladder.
   7. Place a small roll of lint-free tissue in the abdomen and behind the bladder to help stabilize and elevate it.
   8. Prepare to insert the flared end of the PE10 catheter into the bladder.
      1. In the non-dominant hand, hold the dome of the bladder with Dumont #7 curved microforceps and maintain this grip until the catheter is placed in the bladder.
      2. Use a 21-gauge needle to make a cystotomy in the apex of the dome. Gently probe the cystotomy with a closed pair of #5 curved microforceps to make sure the catheter can easily pass through the hole.
      3. While still holding the bladder dome in the non-dominant hand, place the flared end of the PE10 catheter into the bladder (push the flare down to the bladder neck so that it does not slip out while securing it).
      4. Tie the 6-0 monofilament suture around the dome of the bladder and tubing with the tie placed anterior to the tubing. Be sure to tie the suture as high up on the bladder as possible to avoid artificially reducing bladder capacity. (Figure 4)
      5. Alternatively, secure the catheter using a purse string suture as follows. Make a loose purse string suture on the dome of the bladder using 6-0 monofilament. Follow steps 1.3.8.1 - 1.3.8.3 to perform the cystotomy and insert the catheter. Secure the tube by tying the purse string suture. (Figure 5)
   9. Test the patency and seal of the tube in the bladder by attaching a 0.5 mL insulin syringe with a 30-gauge needle to the distal end of the tube. Slowly fill the bladder with 0.1 - 0.2 mL of 0.9% NaCl until a drop appears at the urethral orifice, then empty the bladder by aspiration. It is important that the bladder can be both filled and emptied.
   10. If no leaks occur at the dome, brace the bladder with a pair of curved microforceps and gently pull on the tubing until the flare is resting against the inside of the bladder dome.
   11. Before closing, remove the small roll of tissue, and make sure that the bladder is in its normal position.
   12. Close the abdominal wall in two layers (muscle and skin) with 6-0 running suture. It is preferable to approximate the rectus abdominis muscle by suturing only the edges of the anterior abdominal fascia (anterior wall of the rectus sheath).
   13. To secure the tubing at the animals back, gently rotate the animal onto its abdomen. Insert the subcutaneous portion of the metal anchor into the interscapular incision. (Figure 12) Use a 6-0 suture to secure the tube and anchor by encircling them with a vertical mattress suture.
   14. Make sure a glue bubble remains above and below the skin to prevent the tube from pulling out. Cut the tube approximately 2 cm above the skin.
   15. Gently insert the 30-gauge plug (step 1.1.5) into the end of the tube to prevent urine from leaking out.
4. Inject 0.5 mL 0.9% NaCl subcutaneously for hydration. Give post-operative analgesia immediately following surgery and maintain for 48 h.
   1. Place the animal back into its cage located under an infrared lamp. Maintain constant observation until the animal moves around the cage freely.
5. Monitor the animal daily and allow it to recover for 5 days before recording.

2. Awake Cystometry Recording

1. Preparation of the recording program, pressure transducer, and infusion pump.
   1. Before anesthetizing the animal, connect the infusion pump, pressure transducer, and 22 G swivel using PE50 tubing. (Figure 6)
   2. Open the recording program (see table of materials for an example), on a computer to calibrate the system pressure and prepare for recording. Make sure to use the same settings during calibration and recording.
      1. Fill a 20 mL syringe with 10 - 15 mL of room temperature 0.9% NaCl and load into the infusion pump. Program the pump to infuse at a rate of 0.6 mL/h.
      2. Secure the pressure transducer at the same height as the animal's bladder or the bottom of the recording cage.
      3. Attach the 22-gauge swivel to the end of the pressure transducer (PE50 - tubing - pressure transducer to swivel)
         NOTE: The swivel is used to prevent the tube from twisting or kinking as the animal moves.
      4. Advance the syringe pump to flush 0.9% NaCl through the system. Make sure to remove all air bubbles before calibrating.
      5. With the recording program running, use a ruler to calibrate the pressure (cm/H₂O). Slowly move the end of the PE50 tether from 0 to 30 cm. Adjust the zero if needed.
         NOTE: The 0-cm mark should be at the same height as the floor of the recording cage and pressure transducer.
   3. Suspend the 22-gauge swivel above the center of the recording cage. Ensure that the cage bottom allows urine to fall onto the collecting device of the balance positioned below the cage. Adjust the height of the tether so the mouse can freely move around the cage without straining or stretching the tubing. (Figure 7)
   4. When finished, check to make sure the system and external PE50 tubing are full of 0.9% NaCl and all air bubbles have been removed.
2. Preparation of the animal for recording
   1. Anesthetize the animal with 2% inhaled isoflurane and place it on its abdomen. Remove the 30-gauge plug and slide the PE10 tubing (bladder catheter) into the end of the PE50 tether. Use hot glue to form a watertight seal.
   2. Turn off the anesthesia and place the animal in the recording cagewith a parallel-wire floor, which will allow urine to fall directly onto a collecting device placed on top of an analytical balance. (Figure 7)
   3. Start the recording once the animal is in the cage, but do not start infusing. Monitor the animal until it fully recovers from the anesthesia. Once the bladder pressure stabilizes, begin infusing 0.9% NaCl at a rate of 0.6 mL/h.
      NOTE: Make a note in the recording program when any changes are made. It is important to have a record of when infusion starts, stops, or irregularities occur.
   4. Check the system for leaks and make sure the animal has easy access to food and water.
   5. Continue recording in a quiet room until three reproducible micturition cycles are obtained.
      NOTE: The animal should be completely undisturbed throughout the recording. Preferably, use remote video monitoring to observe behavior.

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Results

There was no significant difference between the tubing materials and diameters in the consistency of pressure rise and fall within the system during tube occlusion. Bladder wall swelling post intravesical tube implantation was significant for both polyethylene (PE) and polyurethane (PU) materials. On day 2, severe submucosal swelling developed. It occupied half the cross-section of the bladder, leading to obstruction of the lumen. On day 5, the edema resolved completely, leaving the submu...

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Discussion

Optimal material and size of intravesical tubing

To determine the effect tubing diameter has on pressure recordings, we tested different microfluidic tubes; PE50 (0.58 mm ID), polyurethane PU027 (0.4 mm ID), PE25 (0.46 mm ID), and PE10 (0.28 mm ID). For each tube, pressure was recorded with the infusion pump running at 1 mL/h, while rapidly moving the tube vertically from 0 to 30 cm. Initial in vivo experiments attempted to use PE50 tubing, but were unsuccessful due to the size of tubing compared...

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Materials

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