Urinary Bladder Distention Evoked Visceromotor Responses as a Model for Bladder Pain in Mice

The overall goal of this procedure is to establish a physiological model of noxious bladder pain and mice during the entire procedure. Body temperature is maintained with a heating pad and overhead radiant heater. The first step in the surgery is to insert a catheter into the urethra of the anesthetized mouse.

The second step is to make an incision to expose the left abdominal muscle. The third step is to take a bite of muscle using a 21 gauge needle. A bent silver wire electrode is then fed into the needle.

The needle is then pulled back through the muscle, leaving the wire in the muscle. This process is repeated for a second silver wire electrode. The fourth step is to place a ground wire into the mouse's chest.

The fifth step now that the surgery has been completed is to distend the bladder and record the EMG responses. This is done by first connecting the three silver wire electrodes to an amplifier and digitizer to record the viscera motor response or VMR. Next, the bladder catheter is connected to a time pressure regulator and air tank to deliver specific stimuli that distend the bladder with different pressures at 15 millimeters of mercury.

No VMR is recorded. However, at 30 45 and 60 millimeters of mercury, the VMR increases with increasing pressure. When plotted, you can see a graded stepwise increase in the normalized VMR with increasing pressures.

This procedure can help inform one's understanding of visceral bladder pain. In particular, using the urinary bladder distension model, one can investigate the entire nervous system in the context of noxious visceral stimulation. This includes recording and investigating from the peripheral nerves themselves, secondary neurons in the spinal cord, and higher neurons in the brainstem and brain.

Additionally, novel therapeutics for the treatment of bladder pain in humans can be tested in this physiological model of bladder pain. Induce an anesthetic State by placing the mouse in an induction chamber at two to 3%isof fluorine. Remove the mouse from the induction chamber after it has lost its writing reflex.

Place the mouse into the nose cone at two to 2.5%isof fluorine. Next, turn the mouse over so that its dorsal side is facing down. Once turned over, test the depth of anesthesia by pinching toe with forceps.

There should be no response from the animal. Next, gently hold the urethral opening perpendicular to the mouse's body with forceps and insert the tip of the catheter into the urethra. Lower the catheter so that it's parallel to the mouse's body, and gently insert the catheter into the mouse's body.

Gently press down on the abdomen to expel any urine in the bladder. Wick this up with a piece of paper towel. Apply Alcohol to the abdomen to disinfect the area.

Make a one to two centimeter incision in the sterilized area to expose left abdominal muscle. Use the scissors to spread the skin and loosen the fascia so that the superior oblique abdominal muscle is exposed. In this image, the abdominal muscle is near the top of the surgical opening and the lug muscle is at the bottom.

Use a 21 gauge needle to take a small bite of muscle. Push the needle through the muscle. It is important to avoid any blood vessels during this process.

Next, feed one end of the silver wire into the needle. Draw the needle back through the muscle to pull the silver wire into the muscle, the wire should be flush with the bite of muscle. Repeat this process.

Implant a second silver wire, approximately 0.5 centimeters from the first Wire. Next, implant a third silver wire into the chest. This wire acts as the ground.

Apply warm mineral oil to the exposed abdominal muscles to keep them moist. Reposition the mouse so that it is on its side in a very comfortable position. Next, connect the electrode clips to the silver wire electrodes.

Position the three clips so that the wires do not touch. Lower the anesthesia to 1.5%After the surgery is complete. Next, lower the anesthesia level by 0.125%every 15 minutes until the mouse exhibits a flexion response to toe.

Pinch without Vocalization or ambulation. Connect the bladder catheter to the Air tubing that is connected to the time pressure regulator. This is used to control the length of the pre pressure interval, the pressure distension trial, and the post pressure inter trial interval.

Perform a distension at 60 millimeters of mercury to test the mouse's response to distension. A strong EMG signal in excess of 0.5 volts should be seen. If this is seen, perform two more distentions to confirm the response.

If the appropriate response is not seen, you must lower the isof fluorine level. If the mouse begins to vocalize or become ambulatory during the distention trial, you must raise the Isof fluorine level. In the current Example, you can see a strong EMG response.

This would be considered a good bimo response to bladder distension and no change in Isof fluorine level would be necessary. Many studies Utilize graded distentions that begin at low pressures, such as 15 millimeters of mercury and workup to more noxious pressures. Now the data has been collected.

It must be processed and analyzed. The first step is to rectify the entire data set by taking the absolute value of each point. This is most easily accomplished using data analysis software, but can also be done manually.

Now that you have the data rectified, determine the average EMG response during the background section of the experiment. Subtract this background EMG response from each data point in the experiment to obtain the background corrected data set. Next, determine the area under the curve for the pre pressure interval, followed by the area under the curve for a pressure distention trial.

This process is repeated for all pre pressure intervals and pressure distention trials. Here's a screenshot from a set of analyzed data. The graphs are in order from top to bottom, the raw EMG, the pressure recorded from a pressure transducer, the stimulus signal, and finally the area under the curve graph.

In this example, to determine the non normalized VMR for the first pressure distension trial, one must simply subtract the calculated area under the curve at the start of the trial from the area under the curve at the end of the trial. Finally, each pressure distention trial area under the curve is normalized to the lowest measured pre pressure interval trial. This is done by dividing each pressure distention trial by the lowest pre pressure interval trial to get the normalized VMR in volts by seconds.

Here's a graph showing the normalized VMR for a single set of distentions ranging from 15 to 75 millimeters of mercury. You can see a graded increase in the VMR on the Y axis with increasing pressures on the x axis. Furthermore, over three sets of graded distentions ranging from 15 to 75 millimeters of mercury, you can see that the overall VMR from this mouse is consistent between sets.

The stability of this technique allows one to do manipulations that can last multiple hours. Finally, illustrating the importance of careful monitoring a body temperature during the bladder distension VMR procedure. Here are a set of data showing that a decrease in body temperature from 37.5 to 33.5 degrees Celsius leads to a significant decrease in the VMR to 60 millimeters of mercury bladder distension.

After watching this video, you should have a good idea of how to surgically implant wire electrodes and a urinary bladder catheter for use with the urinary bladder distension model of visceral pain. You should also have a better understanding of how data generated by this assay are analyzed. Thanks for watching and good luck with your experiments.