The objective of the present article is to provide a detailed description of the recommended procedures to evaluate respiratory function in conscious mice by double-chamber plethysmography.
This technique will help address key question in various fields including respiratory research, safety pharmacology, and drug development. The main advantage of this technique is that it provides read-outs that can be trusted to assess respiratory function in conscious animal both at baseline and during challenges. The technique is especially useful when the experimental protocol requires little deviation from animals'normal physiological state.
The animal is restrained, but otherwise breathes normally which allows assessment of the breathing pattern in addition airway obstruction. On the day of the experiment, start an experimental session and load the appropriate configuration file. Enter the experiment and the animal's information.
Once done, click on the run button at the bottom of the window. Proceed with the calibration of the system. Calibrate each site and input signal separately.
Turn on the bias flow generator connected to the head chamber via a piece of tubing and adjust the flow rate. Close the top opening of the head chamber with a cap. Detach the back panel of the thoracic chamber.
Then firmly insert the calibration tool inside the rubber opening between the head and the body chamber to create a hermetic seal. Close and re-attach the back panel of the thoracic chamber. From the software toolbar menu, go to Tuning and then Calibrate.
Go to Input one and select calibrate to launch the calibration dialogue box for the thoracic flow signal. Verify that the listed parameters in the calibration dialogue window display the appropriate settings. The physical stress applied low value should be zero.
The physical stress applied high value should be minus 20 milliliters per second and samples should be set to integrate. Once done, click on low in the samples window. Verify that the signal generated is constant across the display window and then click on close.
Connect a 20 milliliter syringe through the side port of the thoracic chamber using a plastic connector in a piece of tubing. Select high in the samples window and immediately inject 20 milliliters of air into the chamber over a two second period at a flow rate as constant as possible. Verify that the signal generated appears completely inside the display window.
Use the arrow icon to verify whether the signal is centered and symmetric around the zero line. Then click on close. Remove any offset from zero by clicking on Remove AC offset in the samples window.
Calibrate the head chamber in a similar manner as the thoracic chamber by selecting input number two. This time, set the high value to plus 20 milliliters per second instead of minus 20. Work in a quiet area remote from the housing room.
Acclimate the animals to the restrainer and procedures prior to the start of the experiment. Because it is not invasive and does not require anesthesia, working under conditions where the animal is comfortable, well-adapted, and calm will limit the propensity of hair leaks between the chambers and will thereby maximize the quality of the data. Weigh the animals.
Select the appropriate restrainer for the animal. Insert the animal within the restrainer proceeding from the back opening. Holding the device vertically can be helpful.
Once the animal is in position, insert the back plunger and gently lock it in place without applying an excessive force. Visually check that the animal breathes normally. If needed, adjust its position by moving the locking mechanism.
Ensure that the animal's nares are protruding outside of the nose cone with its snout resting against the inner walls of the restrainer. Detach the back panel of the thoracic chamber. Insert the restrainer containing the animal through the rubber opening in the thoracic chamber and close the chamber.
Attach the head chamber, provide a bias flow, and allow the animal to relax for at least five minutes. Once the animal is calm, start the protocol of commands by selecting the first step and then click on Execute. Check on the computer screen that the animal's breathing signals are regular and smooth.
The software automatically displays the calculated parameters on a breathe by breathe basis. Verify that the animal's parameters are stable. Record the breathing pattern under baseline conditions for up to 10 minutes.
For protocols involving the administration of a test substance by aerosol, first, adjust the nebulizer on time and duty cycle as required. Perform a vehicle challenge and record the response. If needed, expose the animal to increasing concentrations of the test substance by changing the concentration in the nebulizer and escalating steps.
Record the response after each administration. In this example 15 milligrams per milliliter of methacholine is tested. The peak increase in specific airway resistance is recorded a moment thereafter and is then seen to slowly return to baseline value.
At the end of the experimental session, return the animal to its room and housing cage. Between sessions, clean the plethysmograph chambers and rinse the nebulizer with water. Proceed to data analysis as described in the text protocol.
Shown here is the outcome of a repeated respiratory function evaluation at baseline over three consecutive days in two groups of BALB/c mice. A control one and one with pulmonary allergic inflammation. Stable and comparable values were obtained for a selection of parameters provided by double chamber plethysmography.
These values were plotted for breathing rate, title volume, minute ventilation, end-inspiratory pause, flow at mid-title expiratory volume, and specific airway resistance. Changes in respiratory function and response to increasing doses of inhaled methacholine were also performed on successive days to assess the degree of responsiveness in control and allergic C57BL/6 mice. The results display the expected progressive increase in specific airway resistance with augmenting doses of methacholine with an exaggerated degree of responsiveness in the allergic mice on the second and on the third day.
On the following day Newtonian resistance was measured with forced oscillation technique. Changes in this parameter mainly reflect variations in resistance of the large conducting airways. The measured Newtonian resistance at baseline and in response to methacholine correlated well with the measured specific airway resistance obtained the previous day in the same animals.
This technique is convenient to assess numerous animals at once, to capture the kinetics of an acute response, as well as to measure respiratory function in a repeated manner during an experiment that lasted several days. Following the procedure, other methods like the forced association technique can be performed in order to complement the evaluation with direct measurements of respiratory mechanics such as airway resistance. The double chamber plethysmography truly provides an interesting approach to evaluate the breathing pattern and the geography of airway obstruction especially when the experimental protocol required the animals to be conscious.
