Reagents
Acetyl-β-methylcholine (methacholine, MCh), cyclophosphamide monohydrate, porcine pancreatic elastase (PPE) and lipopolysaccharide (LPS) were supplied from Sigma-Aldrich (Bornem, Belgium). Formaldehyde (36%) was obtained from VWR international (Leuven, Belgium) and was diluted to 4% in distilled water. Bleomycin sulfate (Bleo) was supplied from Sanofi-Aventis (Diegem, Belgium), Isoflurane (Forene) from Abbott (Ottignies, Belgium), Pentobarbital sodium (Nembutal) from Sanofi Santé animale (CEVA, Brussels, Belgium), Xylazine from VMD S.A. (Arendonk, Belgium) and Ketamine from Eurovet Animal Health (Bladel, Netherlands).
Animals and disease models
Male BALB/c mice, obtained from Harlan (The Netherlands) at 9 weeks old, were divided into 5 different groups: bleomycin (Bleo)-induced lung fibrosis (n = 10), porcine pancreatic elastase (PPE)-induced emphysema (n = 10), lipopolysaccharide (LPS)-induced acute lung injury (ALI) (n = 8), house dust mite (HDM)-induced allergic asthma (n = 7), and an untreated control group (n = 8). Due to expected mortality in the Bleo fibrosis and the PPE emphysema groups, we included two extra mice in each one. All mice were housed in a conventional animal facility with 12-h dark/light cycles. They were housed in filter top cages and received lightly acidified water and pelleted food ad libitum. All experimental procedures performed in mice were approved by the KU Leuven local Ethical Committee for animal experiments (P166-2012).
Fibrosis
To induce pulmonary fibrosis, mice (10 weeks old) received an intraperitoneal injection of cyclophosphamide (100 mg/kg). Two days later, the animals were anesthetized with an intraperitoneal injection of ketamine/xylazine (100 mg/kg ketamine and 10 mg/kg xylazine; 100 μL/10 g) and intratracheally instilled with bleomycin. Bleomycin was dissolved in sterile PBS and administered as a single dose of 0.1 unit/40 μL per animal [11,12,13]. At the age of 14 weeks, i.e. three weeks after the first bleomycin administration, mice were evaluated (see Additional file 1: Figure S1).
Emphysema
Emphysema was induced in mice (9 weeks old) by administration of porcine pancreatic elastase (PPE). PPE was dissolved in sterile saline and administered in a dose of 1.5 units/50 μL per animal by intranasal instillation, after light anesthesia with isoflurane. Mice were subsequently exposed on a weekly basis to 1.5 units PPE via the same administration route during a period of three weeks [11, 12, 14]. At the age of 14 weeks, i.e. three weeks after the last PPE instillation, mice were evaluated (see Additional file 1: Figure S1).
Acute lung injury
To induce acute lung injury, mice (13 weeks old), received a single intranasal instillation of LPS. LPS was dissolved in sterile saline and administered as a single dose of 3 μg/40 μL/mouse after light anesthesia [15]. The mice were studied at the age of 13 weeks, i.e. two days after the instillation (see Additional file 1: Figure S1).
Allergic asthma
BALB/c mice (11 weeks old) were endonasally sensitized with 1 μg of HDM extract (Greer Laboratories, Lenoir, NC) in 50 μL of saline at day 1. At days 8 to 13, mice were endonasally challenged with 10 μg of HDM extract in 50 μL of saline [16]. Two days after the last challenge, i.e. at the age of 13 weeks, lung function measurements were performed (see Additional file 1: Figure S1).
Lung function measurements
FOT and FE measurements were performed using the flexiVent FX system (SCIREQ Inc., Montreal Qc, Canada). The system was equipped with a FX1 module as well as with a NPFE extension for mice and it was operated by the flexiWare v7.2 software. A small particle size Aeroneb Lab nebulizer (2.5–4 μm; Aerogen, Galway, Ireland) was integrated in the inspiratory arm of the Y-tubing for the generation of the aerosol challenges. The nebulizer activation was synchronized with inspiration and set to a 50% duty cycle for 5 s. On the day of the experiment, mice were anesthetized with an IP injection of pentobarbital sodium (70 mg/kg body weight). Once a surgical plane of anaesthesia was reached, the trachea was exposed to insert an 18-gauge metal cannula having a typical resistance of 0.3 cmH2O.s/mL. Mice were quasi-sinusoidally ventilated with a tidal volume of 10 mL/kg, a frequency of 150 breaths/min, an inspiratory to expiratory ratio of 2:3, and a positive end-expiratory pressure of 3 cmH2O.
A. Baseline measurements
At the start of an experiment, two deep inflations were successively applied to maximally inflate the lungs to a pressure of 30 cmH2O in order to open-up closed areas and standardize lung volume. The lungs were allowed to equilibrate at that pressure over a period of 3 s and the gas compression-corrected volume was read as the subject’s inspiratory capacity (IC), typically on the second maneuver. A broadband forced oscillation waveform inducing frequencies between 0.5 to 19.75 Hz (Prime-8; P8) was then applied to the subject’s airway opening during 8 s. This FOT measurement was analyzed by fitting the constant-phase model to the respiratory input impedance calculated from the P8 experimental data [2]. This mathematical model allows the separate assessment of the airway and tissue contributions to the respiratory response. The Newtonian resistance (Rn, airway resistance, which is dominated by the resistance of the large conducting airways), tissue damping (G, closely associated to tissue resistance and the resistance of the small peripheral airways), tissue elastance (H), and tissue hysteresivity (eta) (G/H) were considered in this study, provided that the coefficient of determination of the model fit was ≥ 0.9. This was followed by the construction of a pressure-volume (PV) curve using a ramp-style pressure-driven maneuver (PVr-P), from which the static compliance (Cst) was calculated directly from the deflating arm of the PV loop between the pressures of 3 –7 cmH2O. Considering the high distensibility of emphysematous lungs, PV loops of mice with emphysema were only reported if the maximum inflation pressure of +30 cmH2O was reached. The hysteresis (Area), or area between inflation and deflation limb of the PV curve, was also calculated. Finally, a NPFE maneuver was performed to obtain a FV loop and the FE-related parameters. This was done by first inflating the lungs to a pressure of +30 cmH2O over 1.2 s and then rapidly exposing the subject’s airways to a negative pressure of -55 cmH2O to generate an imposed negative expiratory pressure gradient. From the FE parameters, the forced expiratory volume and flow at 0.1 s (FEV0.1, FEF0.1, respectively), forced vital capacity (FVC), and peak expiratory flows (PEF) were considered in the present study as well as the FEV0.1/FVC (referred to as Tiffeneau index in humans at 1 s).
B. Airway responsiveness assessment
After performing the sequence of measurements at baseline level, a protocol for measuring airway responsiveness (AHR) to methacholine (MCh) was initiated. Each mouse was exposed for 5 s to an aerosol of a solution of MCh in increasing concentrations (0, 1.25, 2.5, 5, 10, 20 mg/mL). Starting almost immediately after the end of the aerosol exposure, an automated sequence of closely-spaced FOT measurements was initiated using the 3 s long, broadband forced oscillation perturbation ‘Quick Prime-3’ (QP3). Five consecutive QP3 perturbations were run at approximately 15 s apart, resulting in 5 measurements for each concentration of MCh. Each sequence was followed by a NPFE measurement taken approximately 15 s after the last FOT measurement. The dose-response curve to MCh was constructed in a cumulative manner with no recruitment maneuvers in between aerosol challenges as this approach had the advantage of enabling the separation between control and disease mice. Since the NPFE-induced derecruitment of lung units was not reversed by a deep lung inflation maneuver in between aerosol challenges and that it was shown to specifically affect the tissue-related parameters G and H [10], Rn was the only FOT-related parameter considered in assessing airway responsiveness to MCh in the present study to avoid any confounding effects.
For each mouse of the LPS-ALI, HDM-asthma and control group, the provocative concentration of MCh inducing either a 20 (PC20), 30 (PC30) or 40% (PC40) decrease in FEV0.1 was assessed, by calculating the slope of the dose-response curve of each individual mouse, where the peak responses to MCh were normalized to the FEV0.1 of 0 mg/ml MCh (=100%).
Inflammatory cells in the bronchoalveolar lavage fluid
Immediately after the lung function measurements, the lungs were lavaged, in situ, three times with 0.7 mL sterile saline (0.9% NaCl), and the recovered fluid was pooled. Cells were counted using a Bürker hemocytometer (total cells) and the bronchoalveolar lavage (BAL) fluid was centrifuged (1000 g, 10 min). For differential cell counts, 250 μL of the resuspended cells (100,000 cells/mL) were spun (1400 g, 6 min) (Cytospin 3, Shandon, TechGen, Zellik, Belgium) onto microscope slides, air dried and stained (Diff-Quik® method, Medical Diagnostics, Düdingen, Germany). For each sample, 200 cells were counted for the number of macrophages, eosinophils, neutrophils and lymphocytes.
Histopathology
The lungs were fixated in 4% of formaldehyde. After dehydration and embedding in paraffin, sagittal sections were stained with hematoxylin and eosin (H&E).
Statistical analyses
For each mouse, the average of all measurements was calculated for Rn, grouped per experimental conditions, and plotted against MCh concentration. The results are presented as mean ± standard deviation. MCh dose-response curves (Rn, FEV0.1, and FVC) were analyzed using a two-way parametric analysis of variance (ANOVA) followed by a Bonferroni multiple comparison post hoc test to compare each treatment group with the control group. The area under the MCh dose-response curve (AUC) was also calculated and statistically analyzed using a one-way parametric ANOVA. All other statistical analyses were performed using unpaired t-tests, to compare each disease entity separately with the control group (GraphPad prism 5.01, Graphpad Software Inc., San Diego, CA).