TITLE: Detection of central and obstructive sleep apneas in mice: a new surgical and recording protocol
AUTHORS: Gabriele Matteoli, Sara Alvente, Chiara Berteotti, Dario Coraci, Viviana Lo Martire, Martina Lops, Elena Miglioranza, Alessandro Silvani, Emilia Volino, Giovanna Zoccoli, Stefano Bastianini*

[DESCRIPTION]
Sleep apnea is a common disorder in humans and consists of recurrent episodes of cessation of breathing or decrease in airflow during sleep. Sleep apneas can be classified as central (CSA) or obstructive (OSA), based on their origin. CSA results from impairments of the inspiratory drive generated by the brainstem with a consequent interruption of diaphragm muscle effort, while OSA occurs due to upper airway obstruction during inspiration with a continued or increased respiratory effort. Mice are valuable tools to expedite the comprehension of the pathophysiology and genetics of sleep apneas and the development of targeted therapies to contrast sleep-disordered breathing and its adverse consequences on health. Therefore, we provide a method for distinguishing between CSA-like and OSA-like events during sleep in mice by simultaneously recording sleep states, breathing pattern, and diaphragm electromyographic activity inside a whole-body plethysmograph.
We applied this technique in adult C57BL/6J wild-type mice and in a mouse model of Down syndrome (PMID: 34509609).

[STEPS]
SECTION: Overview
1. Figure 1 summarizes the chronological steps of the protocol.

Figure 1. Overview of the protocol

The experimental protocol involves the crafting (Day -1) and the surgical implantation (Day 0) of the electrodes for the recording of electroencephalogram (EEG), nuchal muscle electromyogram (nEMG), and diaphragm electromyogram (DIA). 
After an appropriate period of full post-operatory recovery (Day 5), mice underwent a 7-hour session inside a whole-body plethysmograph to simultaneously record EEG, nEMG, DIA, and ventilation (VENT). 
In the analysis phase, EEG and nEMG signals are used for the sleep scoring, while VENT and DIA signals are employed for the classification of sleep apneas.

 

SECTION: Step 1: Crafting of electrodes
2. In this section, we describe how to craft the electrodes for electroencephalogram (EEG), nuchal muscle electromyogram (nEMG), and diaphragm electromyogram (DIA) recordings.


 

SECTION: Step 1: Crafting of electrodes
3. Preparation of EEG electrode (Figures 2A-3A and Video 1)
SECTION: Step 1: Crafting of electrodes
3.1. Cut a pair of 1.2-cm long multi-stranded stainless-steel wires coated with an insulating sheath.
SECTION: Step 1: Crafting of electrodes
3.2. Gently remove 2-3 mm of the coating from both ends of each wire.
SECTION: Step 1: Crafting of electrodes
3.3. Solder one end of each wire to a stainless-steel screw; we suggest using screws with a length of 2.4 mm and a diameter of 1.2 mm.
SECTION: Step 1: Crafting of electrodes
3.4. Solder the uninsulated remaining end of each wire to a connector equipped with two terminal pins (one wire per pin); we suggest using connectors about 0.5 cm wide and 0.3 cm high.
SECTION: Step 1: Crafting of electrodes
4. Preparation of nEMG electrode (Figures 2B-3B and Video 2)
SECTION: Step 1: Crafting of electrodes
4.1. Cut a pair of 4-cm long multi-stranded stainless-steel wires coated with an insulating sheath.
SECTION: Step 1: Crafting of electrodes
4.2. Remove 2-3 mm of the coating from one end of each wire and at 1.5 cm from it.
SECTION: Step 1: Crafting of electrodes
4.3. Solder the uninsulated end of the wire to a connector equipped with two terminal pins (one wire per pin).
SECTION: Step 1: Crafting of electrodes
5. Preparation of DIA electrode (Figures 2C-3C and Video 3)
SECTION: Step 1: Crafting of electrodes
5.1. Cut a pair of 12-cm long multi-stranded stainless-steel wires coated with an insulating sheath.
SECTION: Step 1: Crafting of electrodes
5.2. Bend each wire into a hairpin-like shape.
SECTION: Step 1: Crafting of electrodes
5.3. Twist the wire on itself until it reaches a final length of 6 cm, ensuring the hairpin bend is enlarged and shaped as a circle with about 2 mm in diameter.
SECTION: Step 1: Crafting of electrodes
5.4. Gently melt the coating along the wire using a hot air flux to stabilize the shape of the electrode.
SECTION: Step 1: Crafting of electrodes
5.5. Remove 2-3 mm of the coating from the free end of each wire and the circular hairpin bend.
SECTION: Step 1: Crafting of electrodes
5.6. Solder the uninsulated end of each wire to a connector equipped with two terminal pins (one wire per pin).
SECTION: Step 1: Crafting of electrodes
6. Figure 2. Schematic representation of the electrodes

Schematic representation of electrodes for electroencephalogram (EEG, panel A), nuchal muscle electromyogram (nEMG, panel B), and diaphragm electromyogram (DIA, panel C) recordings.

 

Figure 3. Photographs of the electrodes

Photographs of electrodes for electroencephalogram (EEG, panel A), nuchal muscle electromyogram (nEMG, panel B), and diaphragm electromyogram (DIA, panel C) recordings.

 

Video 1. Crafting of the electroencephalogram electrode

Preparation of the electroencephalogram (EEG) electrode.


 


Video 2. Crafting of the nuchal muscle electromyogram electrode

Preparation of the nuchal muscle electromyogram (nEMG) electrode.


 


Video 3. Crafting of the diaphragm electromyogram electrode

Preparation of the diaphragm electromyogram (DIA) electrode.


 


SECTION: Step 1: Crafting of electrodes
8. 
 

SECTION: Step 2: Sterilization of surgical materials
9. Surgery is performed under aseptic conditions.

Surgical instruments (forceps, tweezers, scissors with a curved tip, mosquito clamps, microsurgical scissors, periosteum remover, and needle holder) must be sterilized by an autoclave cycle at 121 °C for 15 min. Alternatively, they can be disinfected with specific solutions (e.g., 0.35% peracetic acid solutions).
SECTION: Step 3: Surgical implantation of electrodes in mice
10. The procedure is performed under aseptic conditions with the mouse anesthetized (isoflurane 2-2.5% in pure oxygen). 
Analgesic and antibiotic therapy are administered subcutaneously at the beginning and at the end of the surgery, respectively. During the entire duration of the surgery, the mouse is placed over a thermoregulated heating pad to prevent hypothermia.

 

SECTION: Step 3: Surgical implantation of electrodes in mice
11. Anesthesia and surgical field disinfection
SECTION: Step 3: Surgical implantation of electrodes in mice
11.1. Place the mouse in a closed chamber to induce the anesthesia.
SECTION: Step 3: Surgical implantation of electrodes in mice
11.2. Place the mouse on the surgical table, ensuring that its nose and mouth are located within the anesthesia tube.
SECTION: Step 3: Surgical implantation of electrodes in mice
11.3. Shave the mouse in the subcostal area (under the last rib) and the skin above the head between the ears (Figure 4).
SECTION: Step 3: Surgical implantation of electrodes in mice
11.4. Disinfect the shaved surgical fields with an iodine solution.
SECTION: Step 3: Surgical implantation of electrodes in mice
12. Figure 4. Surgical fields

Photographs of the surgical fields after shaving showing the subcostal area (under the last rib, panel A) and the skin above the head between the ears (panel B).

 

SECTION: Step 3: Surgical implantation of electrodes in mice
13. DIA electrode implantation
SECTION: Step 3: Surgical implantation of electrodes in mice
13.1. Place the mouse on an inclined plane with the head at a lower level than the rest of the body in order to obtain a better exposure of the surgical field.
SECTION: Step 3: Surgical implantation of electrodes in mice
13.2. Use scissors to make a 0.5-cm vertical skin incision over the skull between the ears (Figure 5A and Video 4).
SECTION: Step 3: Surgical implantation of electrodes in mice
13.3. Make a second 0.5-cm vertical skin incision on the flank just below the last rib (Figure 5B and Video 4).
SECTION: Step 3: Surgical implantation of electrodes in mice
13.4. Using surgical scissors with curved tips, create a subcutaneous tunnel connecting the two incisions (Video 5).
SECTION: Step 3: Surgical implantation of electrodes in mice
13.5. Pass the DIA electrode subcutaneously from the head to the flank incision with the help of surgical forceps and a mosquito clamp. The electrode connector must remain on the animal’s head (Figure 6 and Video 5).
SECTION: Step 3: Surgical implantation of electrodes in mice
13.6. Place the mouse on the right flank.
SECTION: Step 3: Surgical implantation of electrodes in mice
13.7. With the help of a surgical microscope, make a 0.5-cm horizontal cut in the abdominal muscle wall and the peritoneum under the last rib using microsurgical scissors (Figure 7A).
SECTION: Step 3: Surgical implantation of electrodes in mice
13.8. Identify the last rib attached to the sternum and the eighth intercostal space (i.e., the space between the two last ribs connected to the sternum) (Figure 7B).
SECTION: Step 3: Surgical implantation of electrodes in mice
13.9. Position the DIA electrode in the correct position (Figure 8). Video 6 provides a detailed step-by-step guide for implanting the DIA electrode, illustrating the following process:

Pass a thin non-absorbable surgical thread through the eight intercostal space from outside to inside the abdominal cavity.
Bend the DIA electrode wires to adapt to the natural shape of the abdominal cavity of the mouse.
Pass the surgical thread through one of the circular hairpin bends of the DIA electrode.
Pass the surgical thread back through the same intercostal space in the peritoneal wall, this time proceeding from intrabdominal to external.
Cut the surgical thread, making sure to leave enough length to tie a knot.
Insert the DIA electrode’s circular hairpin bend into the abdominal cavity.
Pull the suture thread until the circular hairpin bend of the DIA electrode is in contact with the abdominal surface of the diaphragmatic muscle and tie a knot.
SECTION: Step 3: Surgical implantation of electrodes in mice
13.10. Repeat the same procedure for the second DIA electrode’s circular hairpin bend, ensuring that the uninsulated circular portions of the electrode are positioned close to each other, but avoiding any contact between them (Video 6).
SECTION: Step 3: Surgical implantation of electrodes in mice
13.11. Apply suture stitches to close the abdominal muscles around the electrode wires. A second series of stitches must be applied to the flank skin incision (Figure 9).
SECTION: Step 3: Surgical implantation of electrodes in mice
14. Figure 5. Skin incisions

Photographs of the vertical incision over the skull between the ears (panel A) and on the flank just below the last rib (panel B).

 

Figure 6. Insertion of the diaphragm electrode

Photographs of the top (panel A) and side (panel B) view of the mouse with the diaphragmatic electrode inserted in the subcutaneous space.

 

Figure 7. Identification of the last rib

Panel A: photograph of the incision in the abdominal muscle wall and the peritoneum under the last rib.
Panel B: magnification of the last rib attached to the sternum (black arrows) and the eighth intercostal space (VIII).

 

Figure 8. Schematic representations of the diaphragmatic electrode positioning

Circular uninsulated hairpin bends of the diaphragmatic (DIA) electrode were sutured to the muscles of the peritoneal cavity in the eighth intercostal space, so that the DIA electrode was in contact with the abdominal surface of the diaphragm (orange curved lines).

 

Figure 9. Surgical sutures

Photographs of the suture stitches applied to close the abdominal muscles around the diaphragm electrode wires (panel A) and the flank skin incision (panel B).

 

Video 4. Skin incisions

Execution of the skin incisions over the skull between the ears and on the flank just below the last rib.


 


Video 5. Insertion of the diaphragm electromyogram electrode

Insertion of the diaphragm electromyogram (DIA) electrode in the subcutaneous space between the skin incisions on the head and the flank.


 


Video 6. Implantation of the diaphragm electromyogram electrode

Detailed step-by-step guide for implanting the diaphragm electromyogram (DIA) electrode.

 

SECTION: Step 3: Surgical implantation of electrodes in mice
15. EEG electrode implantation
SECTION: Step 3: Surgical implantation of electrodes in mice
15.1. Place the mouse in a prone position on the stereotaxic apparatus to immobilize and expose the skull (Figure 10).
SECTION: Step 3: Surgical implantation of electrodes in mice
15.2. Remove the periosteum from the skull and locally apply a drop of hydrogen peroxide (3% H2O2) to remove debris and better highlight the cranial sutures.
SECTION: Step 3: Surgical implantation of electrodes in mice
15.3. Identify bregma and lambda (i.e., the anterior and posterior crossing points of cranial sutures) (Figure 11A and B).
SECTION: Step 3: Surgical implantation of electrodes in mice
15.4. Using a surgical microdrill, make four holes in the frontal and parietal bones 1 mm anterior and 1 mm lateral to both bregma and lambda (Figures 11C and 12A).
SECTION: Step 3: Surgical implantation of electrodes in mice
15.5. Insert two stainless-steel screws in the holes on one side of the skull in contact with the dura mater for better anchoring and balancing the head cap structure to the skull (Figure 12B).
SECTION: Step 3: Surgical implantation of electrodes in mice
15.6. Securely fasten the screws by turning them clockwise (usually one complete turn is sufficient; pay attention not to screw them excessively, which could damage the underlying cerebral cortex).
SECTION: Step 3: Surgical implantation of electrodes in mice
15.7. Insert the two screws of the EEG electrode in the two remaining contralateral holes of the skull in contact with the dura mater (Figure 12C).
SECTION: Step 3: Surgical implantation of electrodes in mice
15.8. Joint the screws and the connector of the EEG electrode to the skull bone using cement for dental use (e.g., biocompatible and atoxic resins composed of methacrylate and apatite).
SECTION: Step 3: Surgical implantation of electrodes in mice
16. Figure 10. Stereotaxic apparatus

Photograph of an immobilized and exposed mouse skull.

 

Figure 11. Mouse's head and screws positioning

Panel A. Photograph of a mouse skull with bregma and lambda (anterior and posterior crossing points of cranial sutures, respectively) marked after the application of hydrogen peroxide.
Panel B: Magnifications of a mouse skull before (left) and after (right) hydrogen peroxide application.
Panel C: Schematic representation of the mouse head and screws positioning. The four holes were made 1 mm anterior and 1 mm lateral to bregma and lambda on both side on the frontal and parietal bones. Electroencephalographic (EEG) electrode screws were inserted in the holes to the right, while the anchoring screws were inserted in the holes to the left.

 

Figure 12. Implantation of the electroencephalographic electrode

Panel A: photograph (left) and magnification (right) of four holes in the frontal and parietal bones performed using a surgical microdrill.
Panel B: insertion of the two anchoring stainless-steel screws in the holes on the left side of the skull.
Panel C: insertion of the screws of the electroencephalographic electrode in the holes on the right side of the skull.

 

SECTION: Step 3: Surgical implantation of electrodes in mice
17. Implantation of the nEMG electrode
SECTION: Step 3: Surgical implantation of electrodes in mice
17.1. Insert the nEMG electrode wires bilaterally in the nuchal muscles using a curved needle, ensuring that the uninsulated portion of the wire is located inside the muscle (Figure 13 A and B).
SECTION: Step 3: Surgical implantation of electrodes in mice
17.2. Block the wires with a knot to keep them in position (Figure 13C).
SECTION: Step 3: Surgical implantation of electrodes in mice
17.3. Incorporate the connectors of the EEG, nEMG, and DIA electrodes into a biocompatible acrylic paste for dental use on the mouse’s skull (Figure 14).
SECTION: Step 3: Surgical implantation of electrodes in mice
18. The implantation of the EEG and nEMG electrodes is also explained in detail in Video 7.

 

Figure 13. Implantation of the nuchal electromyographic electrode

Panels A and B show the insertion of the curved needle on the left and right sides of the neck muscles.
Panel C shows the final position of the two wires of the nuchal electromyographic electrode secured with a knot.Panel C shows the final position of the two wires of the nuchal electromyographic electrode secured with a knot.Panel C shows the final position of the two wires of the nuchal electromyographic electrode secured with a knot.Panel C shows the final position of the two wires of the nuchal electromyographic electrode secured with a knot.

 

Figure 14. Acrylic structure on mouse’s skull

Photographs of the top (panel A) and side (panel B) view of the mouse with the biocompatible acrylic head cap incorporating the electrodes for electroencephalogram (EEG), nuchal muscle electromyogram (nEMG), and diaphragm electromyogram (DIA) recordings.

 

Video 7. Implantation of the electroencephalogram and nuchal muscle electromyogram electrodes

Detailed step-by-step guide for implanting the electroencephalogram (EEG) and nuchal muscle electromyogram (nEMG) electrodes.

 


VIDEO 8

Video 8. Mouse recovery

Typical mouse health condition and behavior the day after the surgery.

 

SECTION: Step 4: Recording within a whole-body plethysmograph
19. In this section, we describe how to prepare and perform the simultaneous recording of the animal’s behavioral states, ventilation, and diaphragmatic activity with whole-body plethysmography (WBP). 

The WBP apparatus consists of two chambers: one designed for housing the mouse, and a separate reference chamber. Each chamber is equipped with air inlet and outlet tubes. The mouse chamber includes temperature and humidity sensors, and a high-precision differential pressure transducer measures the pressure difference between the two chambers, allowing the detection of the animal’s breathing. The mouse body temperature should also be measured ideally.


 

SECTION: Step 4: Recording within a whole-body plethysmograph
20. Preparation of the animal
SECTION: Step 4: Recording within a whole-body plethysmograph
21. Recording within the WBP chamber
SECTION: Step 4: Recording within a whole-body plethysmograph
21.1. Insert the mouse into the WBP chamber (Figures 15 and 16). We suggest equipping the apparatus with a rotating electrical commutator to prevent the twisting of the recording cable (Figure 17).
SECTION: Step 4: Recording within a whole-body plethysmograph
21.3. At the end of the session, after removing the mouse from the chamber, calibrate the WBP system by repeatedly injecting and withdrawing air into the mouse chamber with a calibrated micro-syringe while recordings of the differential pressure between WBP chambers are still running. 

We recommend sequentially injecting and withdrawing 10, 20, 30, 40, and 50 microliters of air at the anticipated mean breathing rate of the mouse. These data will be used to construct a calibration curve to convert WBP pressure deflection into air volumes.
SECTION: Step 4: Recording within a whole-body plethysmograph
22. Figure 15. Whole-body plethysmograph

Schematic representation of the modified 2-chamber whole-body plethysmograph, used in our laboratory. The figure shows the Plexiglas block used to reduce the internal volume of the mouse chamber (A), the mouse chamber with the tower equipped with the rotating electrical swivel (B), and the reference chamber (C).

 

Figure 16. Whole-body plethysmograph

Photographs of the modified 2-chamber whole-body plethysmograph (panels A and B).

 

Figure 17. Top of the whole-body plethysmograph

Panel A shows a schematic representation of the part of the whole-body plethysmograph that was equipped with a rotating electrical commutator (blue-lined segment and cylinders) that prevented the twisting of the recording lightweight cable (green segment) connected to the electrodes included in the acrylic cap on the mouse’s head (red semicircle).
Panel B shows photographs of the top of the whole-body plethysmograph equipped with the rotating electrical commutator.

 


 

 

SECTION: Step 5: Discrimination of wake-sleep states in mice
23. In this section, we describe how to discriminate the different behavioral states in mice based on the EEG and nEMG signal traces according to published criteria.

A more detailed description with figures and a specific MATLAB script for wake-sleep state discrimination in mice can be found in Bastianini et al., 2014 (2). 

 


We recommend scoring mouse sleep with a 4-second time resolution.


 

SECTION: Step 5: Discrimination of wake-sleep states in mice
24. Perform the sleep scoring of the entire recording by visual inspection of raw EEG and nEMG signals (2–4) with the following criteria:

Wakefulness (W) is assigned when the nEMG tone is high and variable, and the EEG is at a low voltage with possible δ (0.5 – 4 Hz) and θ (6 – 9 Hz) frequency components.

Non-rapid-eye movement sleep (NREMS) is scored when the nEMG tone is lower than in W, and the EEG is at a high voltage with prominent δ frequency components.

Rapid-eye movement sleep (REMS) is defined in the presence of muscle atonia (flat nEMG) with occasional muscle twitches, and when the EEG is at a low voltage with predominant θ activity.

Epochs with artifacts or borderline signals should be marked as “undetermined” and excluded from the successive analyses.


 

 

 

SECTION: Step 5: Discrimination of wake-sleep states in mice
25. Figure 18. Discrimination of the wake-sleep states

Example of raw electroencephalographic (EEG) and nuchal electromyographic (nEMG) data obtained during wakefulness (W), non-rapid-eye-movements sleep (NREMS), and rapid-eye-movements sleep (REMS) in mice in our laboratory.

 

SECTION: Step 6: Sleep apnea detection and categorization into CSA and OSA
26. In this section, we first describe how to detect sleep apnea from the WBP signal and then, how to categorize sleep apneas into CSA and OSA based on the joint evaluation of the WBP and DIA signals.

 

SECTION: Step 6: Sleep apnea detection and categorization into CSA and OSA
27. Sleep apnea detection
SECTION: Step 6: Sleep apnea detection and categorization into CSA and OSA
27.1. Using commercially available or custom-made algorithms, calculate the ventilatory period (VP, i.e., the duration of each breath) on the raw WBP tracings, computed as the time period between two consecutive peaks of the WBP signals.
SECTION: Step 6: Sleep apnea detection and categorization into CSA and OSA
27.2. Calculate the mean VP value for each mouse in each sleep state (i.e., NREMS and REMS).
SECTION: Step 6: Sleep apnea detection and categorization into CSA and OSA
27.3. Calculate the apnea threshold for each sleep state, setting it at either two times (2x criterion) or three times (3x criterion, which is the more conservative criterion used in our lab) the mean VP values defined for each mouse.
SECTION: Step 6: Sleep apnea detection and categorization into CSA and OSA
27.4. Using commercially available or custom-made algorithms, for each sleep state, detect all the breaths longer than the apnea threshold and discard false detections due to artifacts (e.g., due to high-amplitude mechanical pressure artifacts caused by the opening or closing of laboratory doors) by visual evaluation of the raw tracings.
SECTION: Step 6: Sleep apnea detection and categorization into CSA and OSA
28. Figure 19 shows a representative tracing of the WBP differential pressure signal recorded inside the WBP chamber in our laboratory.

Figure 19. Ventilatory period

The figure shows a representative tracing of the whole-body plethysmography differential pressure signal (in red) recorded in mice in our laboratory. Grey segments identify the ventilatory period (VP, i.e., the duration of each breath) associated with an apnea (red bracket) and some normal breaths (green bracket). The time scale is one second.

 


 

SECTION: Step 6: Sleep apnea detection and categorization into CSA and OSA
29. Categorization into CSA and OSA of each manually checked apnea episode
SECTION: Step 6: Sleep apnea detection and categorization into CSA and OSA
29.1. Classify the apnea as CSA when the absence of activity in WBP is concomitant with the absence of activity in the DIA signal.
SECTION: Step 6: Sleep apnea detection and categorization into CSA and OSA
29.2. Classify the apnea as OSA when the absence of activity in WBP is coupled to one or more activity bursts on the DIA signal (i.e., with a clear activation of the diaphragm).
SECTION: Step 6: Sleep apnea detection and categorization into CSA and OSA
30. Figures 20 and 21 show representative examples of CSA and OSA events recorded inside the WBP chamber in our laboratory.

Figure 20. Central sleep apneas

Representative examples of central sleep apneas (CSA) recorded inside a whole-body plethysmography chamber in mice in our laboratory. The whole-body plethysmography differential pressure signal (VENT) is represented in red, while the diaphragmatic electrical activity (DIA) is in blue. Green brackets identify CSA events and blue brackets show the lack of contraction of the diaphragm during the apnea.

 

Figure 21. Obstructive sleep apneas

Representative examples of obstructive sleep apneas (OSA) recorded inside a whole-body plethysmography chamber in mice in our laboratory. The whole-body plethysmography differential pressure signal (VENT) is represented in red, while the diaphragmatic electrical activity (DIA) is in blue. Red brackets identify OSA events and blue circles show the contraction of the diaphragm during the apnea.

 


 

SECTION: Step 1: Crafting of electrodes
7. 
 

SECTION: Step 4: Recording within a whole-body plethysmograph
20.1. Before the recording, slightly anesthetize (1–2 minutes under 1-2% isoflurane anesthesia) the mouse to plug a lightweight cable to the electrodes in the head structure for the acquisition of EEG, nEMG, and DIA signals.
SECTION: Step 4: Recording within a whole-body plethysmograph
21.2. The chamber must be continuously purged with a suitable gas mixture to prevent carbon dioxide build-up. The airflow rate should be adjusted according to the size of the plethysmography chamber. For example, our chambers have a 0.97-liter internal volume and are ventilated with breathable air from cylinders at a rate of 1.5 liters per minute.

The animal respiratory signal is measured as the differential pressure between the chamber containing the animal and a second reference chamber through a high-precision differential pressure transducer, corrected based on the temperature and humidity signals, and calibrated based on the data obtained with the procedure detailed at point 21.3. Mouse body temperature information should also be used to correct tidal volume estimates, if available.

The EEG, nEMG, and DIA signals are acquired via a lightweight electrical cable, amplified, filtered, digitalized, and stored together with the WBP differential pressure.