Measuring Breathing Patterns in Mice Using Whole-body Plethysmography

[Abstract] Respiratory dysfunction is among the main cause of severe and fatal pathologies worldwide. The use of effective experimental models and methodologies for the study of the pulmonary pathophysiology is necessary to prevent, control and cure these diseases. Plethysmography, a technique for the assessment of lung function, has been widely applied in mice for the characterization of respiratory physiology. However, classical plethysmography methods present technical limitations such as the use of anesthesia and animal immobilization. Whole-body plethysmography (WBP) avoids these issues providing a non-invasive approach for the assessment of the respiratory function in conscious animals. WBP relies on the recording of pressure changes that are produced by the spontaneous breathing activity of an animal placed inside an airtight chamber. During normal respiration, pressure variation is directly proportional to the respiratory pattern of the animal allowing the measurement of the respiratory rate and tidal volume. These parameters are commonly used to evaluate pulmonary function in different physiological and disease models. In contrast to classical plethysmography methods, WBP technique allows reproducible serial measurements as it avoids animal restraint or the use of anesthesia. These key features rend WBP a suitable approach for longitudinal studies allowing the assessment of progressive respiratory alterations in physiological and pathological conditions. This protocol describes the procedures for the measurement of the breathing patterns in mice using the WBP method, the data analysis and results interpretation.

It is a non-invasive, precise approach that does not required anesthesia, as the mouse is unrestrained and conscious. Therefore, the respiratory parameters obtained with WBP in mice reflect basal physiological values since instrumental restraints and/or anesthesia are not applied (Lim et al., 2014;Quindry et al., 2016). Notably, these experimental conditions are key for the use of WBP in longitudinal studies (Cramer et al., 2015;Flanagan et al., 2019). Moreover, WBP in mice is a well-stablished technique that has been applied for the study of a wide range of biological aspects of the respiratory function providing new insights into neuronal network controlling respiratory rhythm (Crone et al., 2012), sleep-related breathing disorders (Bastianini et al., 2017) or the role of inflammation in the control of breathing (Giannakopoulou et al., 2019).
During WBP, the mouse is placed in an airtight chamber where it can move freely and breathe spontaneously. Inspiration and expiration cycles modify the chamber pressure due to gas compression and expansion within the lungs as well as to changes in temperature and humidity of the air as it enters into the respiratory tract (Lundblad et al., 2002;Lim et al., 2014;Bates, 2017). Breathing frequency and tidal volume can be measured based on the variations in the chamber pressure parameters (period and magnitude) (Lundblad et al., 2002;Adler et al., 2004;Bates, 2017).
Other conventional plethysmography methods require the animal to be immobilized (Double-chamber plethysmography, DCP), or need invasive surgery in an unconscious mouse (Forced oscillation technique, FOT), which does not allow animal recovery. DCP consists of two chambers that separate animal head and thorax in two chambers (Mailhot-Larouche et al., 2018). Habituation is needed prior to acquisition data due to restraint-induced stress. Restraint is well known as a major stressor and it alters respiration parameters (Buynitsky and Mostofsky, 2009), which may confound result interpretation.
Another broadly used approach in lung mechanics is FOT, which requires that the subject's respiratory Note: Silica beads must be maintained dehydrated, when hydrated (color indicator shifts from orange to green, Figure 1C) it is necessary to replace the beads or reactivate them by drying them at 150 °C for 24 h. 5. Introduce study name (press close).
6. Close the configuration file version window that appears. 7. Accept calibration option (always recommended).
8. Activate DC mode for calibration (AC mode is automatically set during recording).
9. Set the high flow value to -10 ml/s. 13. Gently inject 1 ml air volume into the chamber using the connected syringe and 1.5 s after press F11 (Record volume) to adjust high calibration volume. Proper calibration range should be between -35 and +35 ml/s (repeat the process otherwise).
14. Remove the syringe and place back the cap. 15. Click Finish button and proceed to respiration recording. 16. A white light should flash on ready from the adaptive amplifier machine.

Notes:
a. Calibration must be performed every time the equipment is switched on. There is no need to calibrate between each animal/trial in the same day. Calibration can also be initiated through the U option from toolbar at any time.
b. High flow value must be ten times the injected calibration volume (step 13) as a correction factor for FDP analysis. 1. Weigh the mouse right before recording.

Open the plethysmography device and place the animal inside.
Note: It is critical not disturbing the transducer when manipulating the chamber device.
Vibrations can affect the calibration. If this occurs recalibrate the system. Avoid installing the equipment in unstable surfaces or close to high-traffic areas.
3. Close the chamber tightly and make sure the animal moves freely.

Select
Protocol option and then Site 1 choice (the pre-configured protocol will be open).
6. Introduce mouse identification, weight and any other comments of interest. After assessing this step, recording will start, (green flashing window on screen may appear).

7.
Advance the protocol to wait period (3 min) step. After this period the protocol will automatically proceed to the following steps.
Note: During this interval we recommend that the user leaves the workspace/room without disturbing the equipment to avoid interferences with the analysis. However, mouse must be monitored by visual resources or from outside the room through the whole session to record motor activity periods as well as any other behaviors.
8. After recording, return the animal to its cage. Clean and wipe the equipment with 70% Ethanol.
Recording will be automatically saved in the assigned project folder.
9. Record next animal by restart button (start from Step D1) or proceed to data analysis. 10. Turn off all the equipment.  c. In the Column data format section, select General and click finish.
5. Data will appear in the workbook with column headings in the top row.   Table 1: