Invasive and noninvasive methods for studying pulmonary function in mice

Abstract

The widespread use of genetically altered mouse models of experimental asthma has stimulated the development of lung function techniques in vivo to characterize the functional results of genetic manipulations. Here, we describe various classical and recent methods of measuring airway responsiveness in vivo including both invasive methodologies in anesthetized, intubated mice (repetitive/non-repetitive assessment of pulmonary resistance (RL) and dynamic compliance (Cdyn); measurement of low-frequency forced oscillations (LFOT)) and noninvasive technologies in conscious animals (head-out body plethysmography; barometric whole-body plethysmography). Outlined are the technical principles, validation and applications as well as the strengths and weaknesses of each methodology. Reviewed is the current set of invasive and noninvasive methods of measuring murine pulmonary function, with particular emphasis on practical considerations that should be considered when applying them for phenotyping in the laboratory mouse.

Figure 1

Diagram of the plethysmograph used for pulmonary function testing of anesthetized, orotracheally intubated mice. A thermostat-controlled water basin (37°C) built in the plethysmograph chamber ensured a body temperature of 34–35°C as measured by rectal thermometer. Defined aerosol concentrations of methacholine, as measured by an aerosol photometer, were delivered into the airways via the orotracheal tube. For calculation of pulmonary resistance (R_L), transpulmonary pressure (P_TP) was recorded via an esophageal tube, and tidal flow was determined by a pneumotachograph attached directly to the orotracheal tube. PT, pressure transducer. Taken from [10] with permission.

Figure 2

Diagram of the barometric whole-body plethysmograph (taken from [35] with permission). (A) Main chamber containing the animal (B) connected to a pressure transducer (C) which is also connected to the reference chamber (B). (D) Pneumotachograph. Main inlet for aerosol. The bias airflow at 0.2 L/min was discontinued during aerosol challenges.

Figure 3

Schematic drawing of the head-out body plethysmograph. The figure illustrates the attachment of the neck collar (made of dental dam with a central hole of 7–8 mm for a 20–25 g mouse) to the plethysmograph. The adapter is put in the front opening of the plethysmograph and a viscoelastic ring is slipped over the fixed rubber dam at the nose of the plethysmograph thus fixing the collar. The conscious animal is then placed in the glass plethysmograph and attached via the conus to a ventilated head exposure chamber. A moveable glass cylinder built in the screw cap enables atraumatic positioning of the mouse. Volume calibration (1–1.5 ml air) of the plethysmograph (front and back opening sealed) is done before each measurement. Before data collection, mice are allowed to acclimatize for at least about 10 minutes in the body plethysmographs.

Figure 4

Characteristic modifications to the normal breathing pattern in conscious BALB/c mice. A: normal breathing pattern of BALB/c mice breathing room air. B: characteristic pattern of airway obstruction during aerosol challenge with MCh, illustrating the decline in EF₅₀. A and B, top tracings: pneumotachograph airflow signals. A and B, bottom tracings: corresponding integrated VT signal. A horizontal line at zeroflow separates inspiratory (Insp; upward; +) from expiratory (Exp; downward; -) airflow. V, tidal flow. VT, tidal volume. TI, time of inspiration. TE, time of expiration. Figure taken from [49] with permission.