Evaluating the Procedure for Performing Awake Cystometry in a Mouse Model

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.

Transcript

The overall goal of this microsurgical procedure is to implant an intravesical tube into the urinary bladder and to record the lower urinary tract function in an awake mouse. This method can help in the acquisition of reproducible experimental data from awake cystometry and the testing of bladder function in healthy and diseased mouse models. The main goal of this technique is to provide a standardized method based on experimental data for performing filling cystometry in a mouse model.

To prepare the tube for implantation, first cut a seven centimeter piece of pe10 tubing, and slowly advance one end toward an open flame to create a flare, quickly withdrawing the tube as soon as the flare develops. Then use the low heat setting on a glue gun to apply three drops of all purpose hot glue at 4.5, 5, and 5.5 centimeters from the flared end of the tube. Use scissors to trim irregularities from the glue bubbles.

Apply hot glue to one end of a 30 gauge needle, which will be used to plug the pe10 tubing after it is implanted. Next confirm the lack of response to toe pinch and apply ointment to the animal's eyes. After sterilizing the upper back and lower abdomen with alcohol and betadine, make a 1.5 centimeter incision between the scalpula, and place the animal in the supine position on top of a 37 degree celsius heating pad covered with sterile drapes under a dissecting microscope.

Now make a 1.5 centimeter lower midline abdominal skin incision. Followed by a matching incision through the fascia along the linea alba and muscle to expose the dome and upper half of the urinary bladder. Keeping the abdominal viscera moist with warm physiological saline, rotate the animal onto it's side to access the incision on the nape of the neck.

Push a narrow hemostat subcutaneously through the incision to start a subcutaneous channel on the back and continuing along the side of the animal. Once the tip of the instrument reaches the bottom of the ribcage, turn the tip towards the midline in the inside of the abdomen and continue advancing the hemostat until the tip is exposed at the abdominal incision under the muscular layer. Once the hemostat tip is inside the incision, pinch the skin above the shoulder and apply traction to help stabilize the animal until the instrument tip is seen at the abdominal incision.

Now use the hemostat to grasp the non flared end of the tubing and slowly retract the hemostat, pulling the end of the tube out of the incision at the back of the neck and adjusting the flared end of the tubing so that it lies directly above the dome of the bladder. Stabilize the bladder with a small roll of lint free tissue. Place a loosely tied, not absorbable 6-O monofilament suture on top of the bladder dome.

Using the non dominant hand and the Dumont number seven curved micro forceps, grasp the dome of the bladder and use a 21 gauge needle in the dominant hand to make a cystotomy in the dome apex. Gently probe the cystotomy with a closed pair of number five curved micro forceps to make sure the catheter can easily pass through the hole and place the flared end of the pe10 catheter into the bladder, pushing the flare down to the bladder neck so that it does not slip out while being secured. With the monofilament tie placed anterior to the tubing, tighten the 6-O suture around the dome of the bladder and tubing as high up on the bladder as possible to avoid artificially reducing the bladder capacity.

Alternatively, place a loose purse-string suture on the dome of the bladder, make a cystotomy as just demonstrated, and insert the flared end of the catheter into the puncture. Then tie the suture around the dome and the tubing as demonstrated for the flat suture. In either case take care to secure the tubing without causing extensive injury to the bladder wall.

If too much tissue is taken, the bladder capacity will be reduced. If too little is taken, the connection will leak. To test the seal and position of the tube in the bladder, attach a 0.5 milliliter insulin syringe equipped with a 30 gauge needle to the distal end of the tube and slowly fill the bladder with 0.1 to 0.2 milliliters of 0.9%sodium chloride until a drop appears at the urethral orifice.

If the seal is incomplete, loop suture tails around the bladder and retie. Then empty the bladder by aspiration. If no leaks occur at the dome, brace the bladder with a pair of curved micro forceps and gently pull on the tubing until the flare is resting against the inside of the bladder dome.

Retest the seal using the previously described method. Trim the tails of the suture, remove the tissue roll, and return the bladder to it's normal position. Then use a running 6-O suture to close the abdominal wall in two layers.

And gently rotate the animal onto it's abdomen. Insert the subcutaneous portion of the metal anchor into the intrascopular incision, and use a 6-O monofilament to secure the tube and anchor with a vertical mattress suture. Confirm that a glue bubble remains above and below the skin to prevent the tube from pulling out and cut the tube approximately two centimeters above the skin.

Then gently insert a 30 gauge plug into the end of the tube to prevent urine from leaking out. Before performing the reading, use pe50 tubing to connect the infusion pump, pressure transducer, and 22 gauge swivel. Then place the anesthetized experimental animal in the prone position and remove the 30 gauge plug.

Connect the tether to the anchor, slide the bladder catheter into the end of the pe50, and use hot glue to form a water tight seal. Place the mouse onto the wire floor. And begin the recording.

Then when the animal recovers from the anesthesia and the bladder pressure stabilizes, begin infusing 0.9%sodium chloride at 0.6 milliliters per hour rate. Typically, on day two after catheter insertion, severe submucosal swelling that occupies half of the cross section of the bladder develops leading to obstruction of the lumen. On day five the edema resolves completely leaving the submucosal areas infiltrated with inflammatory cells that partially invaded the muscularis.

The greatest extent of the tissue swelling observed on days two and day three correlates with the corresponding behavioral voiding with the animals demonstrating a significantly impaired bladder function at these time points. As expected from the histologic data, the voiding frequency normalizes by post operative day five. Intravesical pressure in an awake freely moving animal with minimal movement artifacts is characterized by a baseline pressure of 10 to 15 centimeters of water which may remain unchanged or may gradually increase by no more than 10 centimeters of water during the filling cycle.

Followed by a sudden pulse style pressure increase and drop during voiding. Once mastered, this technique can be completed in 30 minutes if it is performed properly. While attempting this procedure it is important to remember to minimize the damage to the bladder to preserve the physiology of the tissue.

Following this procedure, other methods, like single cycle filling cystometry can be performed to answer additional questions about the functional bladder capacity. After watching this video, you should have a good understanding of how to microsurgically implant a tube into the urinary bladder and to use awake filling cystometey to evaluate lower urinary tract function in mice.

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Journal (Medicine)

Journal (Medicine)

Evaluating the Procedure for Performing Awake Cystometry in a Mouse Model

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