Measuring Respiratory Function in Mice Using Unrestrained Whole-body Plethysmography

The overall goal of this procedure is to measure respiratory parameters from a conscious rodent using unrestrained whole body plethysmography. This is accomplished by first calibrating the bridge amplifier. Once calibrated, a mouse is placed inside the plethysmography chamber and allowed to acclimatize before testing.

Next, the pressure changes that occur within the chamber are recorded. Multiple respiratory parameters are then calculated from the information obtained while using the plethysmography system. Results show that unrestrained whole body plethysmography is a reliable assessment of lung function in rodents.

The main advantages of this technique over existing methods like invasive plethysmography are that the animals are not anesthetized, which means that we're not associating artifacts with anesthesia. In addition, due to the non-invasive nature of this protocol, you, it allows for longitudinal measures. In general, individuals who are new to this technique tend to underestimate the time needed to acclimatize the animals to the plasm chamber.

To begin, set up the data acquisition machine bridge amplifier and pressure transducer. When ready, move to the computer and open the analysis software. Open the channel settings option found in the setup toolbar and change the number of channels being recorded to one.

Next, set up the water column apparatus as demonstrated. Connect two five milliliter serological pipettes with plastic tubing. Fill the columns with water and ensure the water levels are balanced.

With a ruler connect one piece of plastic tubing to the top of each pipette. To calibrate the bridge amplifier, first, attach a one milliliter syringe filled with 300 microliters of air to the right side of the water column. Attach the left side tubing to the connector on the pressure transducer.

Next, open the scroll down menu and select bridge amp. Enter the correct settings and click zero to set the trace at approximately zero millivolts. When ready, depress the one milliliter syringe, the resulting pressure will move the water in the column by one centimeter and show a sudden spike on the software due to the pressure change.

Next, select input units from the bridge amp window and highlight the background trace prior to the spike, otherwise known as the zero region. Click the arrow next to 0.1 to produce the background signal within the ranges of negative 0.002 millivolts and positive 0.002 millivolts. Type zero in the window adjacent to the background signal window.

Highlight the region of increased pressure on the graph from when the syringe was depressed. Click the arrow next to 0.2. The value should be in the range of 0.9 to 1.2 millivolts.

Next, type one in the window. Next to the increased pressure window, go to define units and select centimeters H2O and hit okay. Return to the bridge amp menu.

Select one millivolt and set amplifier to zero. This will complete calibration and the water column can safely be removed. For the best results, test animals should be introduced to the plethysmography chamber one week before the start of experimentation.

To begin, place the temperature relative humidity, probe on the one whole end of the plethysmography chamber and record the temperature, humidity, and barometric pressure. After recording the animal's weight and body temperature, place it in the plethysmography chamber and close the chamber. Next, insert the transducer and syringe in the two holes on the other side of the chamber.

Press start on the software program and record for up to 45 seconds. Make sure to record five seconds of data when the animal is not moving. Stop recording after 45 seconds and remove the mouse from the plethysmography chamber.

Immediately record the chamber temperature and humidity. Return the mouse to its cage spray and wipe the chamber with 80%volume per volume. Ethanol allow the chamber to dry and return to baseline temperature and humidity before proceeding on to the next mouse.

To begin, open the program in full screen and select the data. Open the mini data pad window and select cycle measurements in the left hand column and average cyclic height. In the right hand column of column one, select option, and set the scale for minimum peak detection to one.

This will present the pressure deflation due to tidal volume PT measurement in the mini data pad. Select cycle measurements followed by event count to present the frequency F measurement. Next, select cycle measurements followed by average cyclic period to present the total breathing cycle time T tote seconds measurement to generate peak inspiration and expiration time values, ensure the cursor is directly over the maximum of the peak trough, and add a comment on nine sequential peaks and troughs.

Open a new window and click selection information in the left hand column and duration in the right hand column before selecting. Okay, select the macro found at the top of the program to start recording. Next, locate the first comment box by using the fine tool.

Go to the start of the file, followed by the first comment and select the add to data pad command. Finally, select begin Repeat from the macro commands and confirm that the repeat count window is set at nine. Use the find next option and select the add to data pad command.

When finished, select and repeat. Stop the recording and save the macro based on the animal number. The macro can now be run to obtain the inspiration ti and expiration te time between each comment.

Since expiration and inspiration occurs consecutively, the data appear in this order and needs to be manually split. This breathing trace illustrates appropriate consistent data from a control animal. Nine consecutive comments are added at the peaks and troughs of breathing oscillations to obtain respiratory parameters.

These examples show the most common suboptimal traces that should never be used for analysis. This trace was recorded while the animal was sniffing and moving a trace with oscillations that gradually increase over time is usually caused by condensation and humidity buildup. The noise in this trace is the result of the animal or researcher engaging with the equipment.

This figure demonstrates the lung function of a mouse with bleomycin induced pulmonary fibrosis. Notice the irregular pattern in the breathing trace. Once the technique is mastered and the equipment calibrated, the data can be collected from each animal within five minutes.

The collected data can then be bat analyzed and exported onto an Excel spreadsheet. While attempting this procedure is important to acclimatize the mouse to the plethysmography chamber before collecting data, this may vary between strains of mice. On average, we use adolescent's C 57 black six mice, and this will take four to five days prior to collecting data.