Devos et al. Respiratory Research (2017) 18:123
DOI 10.1186/s12931-017-0610-1

RESEARCH

Open Access

Forced expiration measurements in mousemodels of obstructive and restrictive lungdiseases
Fien C. Devos1, André Maaske2, Annette Robichaud3, Lore Pollaris1, Sven Seys4, Carolina Aznar Lopez5,
Erik Verbeken6, Matthias Tenbusch2,7, Rik Lories5, Benoit Nemery1, Peter HM Hoet1 and Jeroen AJ Vanoirbeek1*

Abstract
Background: Pulmonary function measurements are important when studying respiratory disease models. Bothresistance and compliance have been used to assess lung function in mice. Yet, it is not always clear how theseparameters relate to forced expiration (FE)-related parameters, most commonly used in humans. We aimed tocharacterize FE measurements in four well-established mouse models of lung diseases.
Method: Detailed respiratory mechanics and FE measurements were assessed concurrently in Balb/c mice, using
the forced oscillation and negative pressure-driven forced expiration techniques, respectively. Measurements were
performed at baseline and following increasing methacholine challenges in control Balb/c mice as well as in four
disease models: bleomycin-induced fibrosis, elastase-induced emphysema, LPS-induced acute lung injury and house
dust mite-induced asthma.
Results: Respiratory mechanics parameters (airway resistance, tissue damping and tissue elastance) confirmed
disease-specific phenotypes either at baseline or following methacholine challenge. Similarly, lung function defects
could be detected in each disease model by at least one FE-related parameter (FEV0.1, FEF0.1, FVC, FEV0.1/FVC ratio
and PEF) at baseline or during the methacholine provocation assay.
Conclusions: FE-derived outcomes in four mouse disease models behaved similarly to changes found in humanspirometry. Routine combined lung function assessments could increase the translational utility of mouse models.
Keywords: Fibrosis, Emphysema, Asthma, Acute lung injury, Mice, Forced oscillations technique, Negative pressure
forced expiration, Airway resistance, Tissue elastance, Forced expiratory volume, Forced vital capacity

BackgroundAt the preclinical level in experimental animals, pulmonary
function is generally assessed using the forced oscillation
technique (FOT). This is an invasive method allowing
accurate measurements of physiologically relevant parameters describing the mechanical properties of the respiratory
system. The classical parameters of respiratory mechanics,
resistance and compliance, relate to the opposition to air
moving in and out of the lungs and to the distensibility of
the respiratory system, respectively. These measurements
are based on the analysis of pressure, volume, and flow
signals acquired in response to oscillatory airflow

waveforms applied at the subject’s airway opening [1].
Through the use of complex oscillatory signals andadvanced mathematical model fitting, this technique
is also able to distinguish between airway and tissue
mechanics [2].
FOT measurements have already been used in mousemodels of respiratory diseases to investigate pathophysiological changes associated with fibrosis [3], emphysema [4], or asthma [5–8]. Yet, from a pre-clinicalperspective, the translational value of the mouse as amodel to study respiratory diseases has been challenged
and it is not always clear how changes in respiratory mechanics parameters relate to the commonly measured* Correspondence: jeroen.vanoirbeek@kuleuven.be1
Center for Environment and Health, KU Leuven, Leuven, Belgium
Full list of author information is available at the end of the article© The Author(s). 2017 Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0
International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, andreproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to
the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver(http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Devos et al. Respiratory Research (2017) 18:123

forced expiration (FE) parameters in humans [9].
Nowadays, spirometric-like measures can be performedin mice by rapidly exposing the airways to a negative
pressure in order to generate a forced expiratory flow
signal [10]. The technique is referred to as negative
pressure-driven forced expiration (NPFE) and it can be
performed concurrently with FOT measurements, inthe same animal using a single device, to obtain, in
addition to respiratory mechanics’ parameters,
endpoints resembling the clinically used ones, such as
forced expiratory volume at 0.1 s (FEV0.1), forced vital
capactity (FVC), “Tiffeneau” index (FEV0.1/FVC) or
peak expiratory flow (PEF). As is the case for humansubjects, both FOT and FE outcomes can also bemeasured during a bronchoprovocation test to
increasing concentrations of methacholine to determine airway hyperresponsiveness (AHR) [9, 10].
Previously, Vanoirbeek et al. [11] reported flowvolume (FV) loops and associated FE parameters
measured at baseline in mouse models of asthma, fibrosis and emphysema. In that particular study, FE
measurements were performed with a system
deprived of any aerosol generation or FOT measurement capabilities. Therefore, in order to be able toassess airway responsiveness to aerosolized methacholine, the subjects had to be moved from the FOT
device to the FE system during the course of theexperiment. In addition to the technical challenges
and a loss in precision in the timing of the measurements, the use of that FE system was also associatedwith the need to manually construct the FV curves,which were generated from a total of 9 data points
acquired during the FE maneuver. More recent FE
systems now have the ability to automaticallygenerate FV loops and derive FE parameters. In
addition, the FV curves are constructed from higher
resolution volume and flow signals (containing
approximately 360 data points), thus providing amore accurate and specific assessment of the respiratory system under conditions of greater expiratory
pressure gradient. The objective of this study was toevaluate FE endpoints (FEV0.1, FVC, PEF, FEV0.1/
FVC) as potential diagnostic parameters in different
mouse models of respiratory diseases. In order to doso, we seamlessly performed FOT and NPFE measurements at baseline and after increasing concentrations of methacholine using a single measurement
device that provided an accurate FE assessment.
Since the hypothesis was that FE-related changes
would generally behave similarly to changes found in
human spirometry across a range of respiratory
diseases, we included a model of lung fibrosis,
emphysema, and acute lung injury, as well as amodel of allergic asthma. All four mouse respiratory

Page 2 of 14

disease models were compared to a group of naive
control animals.

Methods
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 fromVMD S.A. (Arendonk, Belgium) and Ketamine from
Eurovet Animal Health (Bladel, Netherlands).
Animals and disease models

Male BALB/c mice, obtained from Harlan (TheNetherlands) 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 lunginjury (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 ina conventional animal facility with 12-h dark/lightcycles. They were housed in filter top cages andreceived lightly acidified water and pelleted food adlibitum. All experimental procedures performed inmice were approved by the KU Leuven local EthicalCommittee 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 asa single dose of 0.1 unit/40 μL per animal [11–13]. At theage 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Devos et al. Respiratory Research (2017) 18:123

anesthesia with isoflurane. Mice were subsequently exposedon 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 sensitizedwith 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 theflexiVent 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 ofthe experiment, mice were anesthetized with an IP
injection of pentobarbital sodium (70 mg/kg bodyweight). Once a surgical plane of anaesthesia wasreached, 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 ventilatedwith 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 weresuccessively applied to maximally inflate the lungs to apressure of 30 cmH2O in order to open-up closed areasand standardize lung volume. The lungs were allowed toequilibrate at that pressure over a period of 3 s and thegas compression-corrected volume was read as the
subject’s inspiratory capacity (IC), typically on the

Page 3 of 14

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 associatedto 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 thecoefficient of determination of the model fit was ≥ 0.9. Thiswas 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 thehigh distensibility of emphysematous lungs, PV loops ofmice 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-relatedparameters. This was done by first inflating the lungs to apressure 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. Eachmouse 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 closelyspaced FOT measurements was initiated using the 3 slong, 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. Eachsequence was followed by a NPFE measurement taken
approximately 15 s after the last FOT measurement. The
dose-response curve to MCh was constructed in acumulative manner with no recruitment maneuvers in
between aerosol challenges as this approach had theDevos et al. Respiratory Research (2017) 18:123

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-relatedparameter considered in assessing airway responsivenessto MCh in the present study to avoid any confounding
effects.
For each mouse of the LPS-ALI, HDM-asthma andcontrol 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 slopeof 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, thelungs 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 μLof 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 thenumber of macrophages, eosinophils, neutrophils andlymphocytes.
Histopathology

The lungs were fixated in 4% of formaldehyde. Afterdehydration and embedding in paraffin, sagittal sections
were stained with hematoxylin and eosin (H&E).
Statistical analyses

For each mouse, the average of all measurements wascalculated for Rn, grouped per experimental conditions,and plotted against MCh concentration. The results arepresented as mean ± standard deviation. MCh doseresponse 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 statisticalanalyses were performed using unpaired t-tests, tocompare each disease entity separately with the control
group (GraphPad prism 5.01, Graphpad Software Inc., San
Diego, CA).

Page 4 of 14

Results
During the induction protocols, two mice of the PPE-treated (emphysema) group and two mouse of the Bleotreated (fibrosis) group died. In both groups, mice had significantly lower body weights on the day of analysis, compared with the control group. In the HDM asthma group,
one mouse was excluded because all data were far out of
the 95th percentile of the group. All data reflect measurements from at least seven mice per group.
Inflammatory cells in BAL fluid

Compared to the control group, significantly higher
numbers of neutrophils were found in the BAL-fluid ofmice with bleomycin-induced fibrosis and LPS-induced
acute lung injury (ALI). Similarly, significantly higher
numbers of eosinophils were found in mice with
HDM-induced asthma, in comparison to naive control
mice. Lymphocytes were significantly increased in micewith bleomycin-induced fibrosis, elastase-induced
emphysema and HDM-induced asthma, compared tonaive controls (Fig. 1a).
Histopathology

To further confirm the underlying pathology, paraffin
sections of the lungs were obtained of every mouse.
Figure 1 shows an illustrative example of each group.
Compared to naive control mice (Fig. 1b), mice withfibrosis presented with a heterogeneous inflammatory
cell infiltrate in both the broncho-vascular bundle andalveolar parenchyma, and fibrous thickening of alveolar
septa (Fig. 1c), whereas mice with emphysema presented
with a typical heterogeneous destruction of the pulmonary parenchyma (Fig. 1d). Mice with ALI showed a mild
infiltration of inflammatory cells, including mononuclear
cells and granulocytes in the broncho-vascular bundle
and intra-alveolar (Fig. 1e). Mice with asthma showed apronounced infiltration of mononuclear cells and granulocytes in the broncho-alveolar bundle and in the venous
vascular tree. Mild intra-alveolar inflammation was alsopresent (Fig. 1f ).
Lung function measurements
A. Baseline measurements

A complete overview of all parameters measured at baseline is reported in the Additional file 2.
FOT respiratory mechanics measurements Consistent
with an anticipated increase in tissue stiffness and elastic
recoil, mice treated with bleomycin displayed, at baseline,
elevated values of G (tissue damping) and H (tissue elastance) relative to naive control mice, with no statisticaldifference in Rn (central airway resistance) (Fig. 2). In contrast, in the PPE-treated mice, where a decrease in elastic
recoil was expected, G and H were significantly decreased

Devos et al. Respiratory Research (2017) 18:123ab

BAL cell count8x104 cells/mL

Page 5 of 14

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Fig. 1 Differential cell counts in bronchoalveolar lavage and histological analyses. Total numbers of macrophages, neutrophils, eosinophils andlymphocytes were determined in the bronchoalveolar lavage (BAL) fluid, obtained immediately after lung function measurements (a). Next, lungs wereisolated and slides of the lungs from mice with different lung diseases were prepared and colored with H&E-staining for analysis at a magnification of
50× and 200×, shown for naive control group (b), Bleo-fibrosis (c), PPE-emphysema (d), LPS-ALI (e) and HDM-asthma (f). Data arepresented as mean ± SD. * p < 0.05, ** p < 0.01 and *** p < 0.001 compared to the naive control group. n = 7 –8 per groupcompared to control animals, resulting in a significantly
increased tissue hysteresivity (G/H). These animals also
showed a significant increase in airway resistance (Rn) incomparison to the control group (Fig. 2a). As seen in
Fig. 1, mice with ALI also presented with an increase inRn, compared to the naive healthy mice. Baseline parameters of mice with HDM-induced asthma did not differ
from those of naive control mice (Fig. 2).
Pressure-volume loop and inspiratory capacity The
PV loop of mice with fibrosis showed a characteristic
downward shift, while those of emphysematous micedisplayed a typical upward shift, compared to naive
control mice (Fig. 3a). Consistent with an increased

stiffness of the lungs, mice with fibrosis showed a significant decrease in IC and Cst compared to naive
controls (Fig. 3c, d). Emphysematous mice, on theother hand, showed a statistically elevated IC with,however, no significant change in Cst relative to thecontrol mice (Fig. 3c). The PV loops from the micewith ALI or with asthma appeared very similar to
those from the naive control ones (Fig. 3b). Consistently, IC and Cst were unchanged across theseexperimental groups (Fig. 3c and d).
FE measurements Forced expiration in mice with fibrosis resulted in a characteristic FV loop compared to control mice, with reduced PEF and FVC suggestive ofDevos et al. Respiratory Research (2017) 18:123

aPage 6 of 14b

Airway resistance (Rn)

Tissue damping (G)
8

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cmH2O.s/mL*

Fig. 2 FOT respiratory mechanics parameters at baseline. Central airway resistance, Rn (a), tissue damping, G (b), tissue elastance, H (c) and tissue
hysteresivity G/H (eta) (d) were measured using a prime-8 perturbation. Parameters are shown for each individual mouse, along with groupmeans. * p < 0.05 and *** p < 0.001 compared to the naive control group. n = 7 –8 per group

restrictive lung disease (Fig. 4a). FV curves of emphysematous mice also displayed a pronounced decrease in
PEF with, however, an increased FVC relative to control
healthy mice (Fig. 4a). The FV curves of mice with ALI
and asthma generally followed the curve of control mice,with some reductions in PEF (Fig. 4b).
These changes in the shape of the FV loops werealso reflected in the FE-derived parameters (Fig. 5).
Consistent with a typical restrictive airway patternseen in human patients, mice with fibrosis showed atbaseline a significantly reduced FEV0.1, FVC and PEF,
as well as a normal FEV0.1/FVC ratio (Fig. 5). In
mice with emphysema FVC and FEF0.1 were significantly increased, while FEV0.1, PEF and FEV0.1/FVC
ratio were significantly decreased. From all FErelated parameters, PEF was significantly decreased
in mice with ALI, compared to naive controls.
HDM-induced asthma mice displayed at baseline FEderived parameters that generally followed what wasseen in the control mice, with the exception of asmall but statistically significant increase in FVC.

B. Assessment of airway responsiveness to methacholine

In this B section, we report MCh responses exclusively formice with LPS-induced ALI and HDM-induced asthma,
as these provide clinically-relevant information.
FOT respiratory mechanics measurements AHR,
expressed in terms of central airway resistance (Rn), wasabsent in mice with ALI (Fig. 6a), but present, as
expected, in mice with asthma (Fig. 6b). In this lattergroup, Rn was significantly increased at the two highest
concentrations of MCh tested, compared to naive
control mice (Fig. 6b).
FE measurements Mice with ALI displayed a similar
FV loop shape profile as the control ones in response
to increasing MCh aerosol challenges (Fig. 7a,b). The
shape of the FV loops of the mice with HDMinduced asthma became more concave with increasing
concentrations of MCh, reflecting a typical pattern ofairflow limitation due to obstruction. The FV curve of
the HDM-treated group also showed a decreasing

Devos et al. Respiratory Research (2017) 18:123bPV-loops baseline

PV-loops baseline

1.2

1.2

0.9

0.9

Volume (mL)

0.6
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Pressure
Naive Ctrls
Bleo-fibrosis
PPE-emphysemacdInspiratory capacity (IC)
***

1.2

20

30

Pressure
Naive Ctrls
LPS-ALI
HDM-asthma

Static compliance (Cst)

0.12

mL/cmH2O

***
mL

0.9

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as o-f-e ibrom
ph sisysemaVolume (mL)aPage 7 of 14

Fig. 3 Pressure-volume curves and related parameters at baseline. Pressure-volume (PV) loops were generated using a ramp-style pressure-volume
maneuver (PVr-P). Mean PV loops are shown for each group, Bleo-fibrosis and PPE-emphysema in (a), LPS-ALI and HDM-asthma in (b), and represented
together with the naive control group. Inspiratory capacity (c) was measured using a deep inflation maneuver, and the static compliance (d) was calculated
directly from the deflating arm of the PV loop between 3 –7 cmH2O. Both parameters are shown for each individual mouse, along with group means. ***p < 0.001 compared to the naive control group. n = 7 –8 per groupb

FV-loops baseline

FV-loops baseline

50

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Flow (mL/s)

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PPE-emphysema

0.0

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Volume (ml)Naive Ctrls

LPS-ALI
HDM-asthma

Fig. 4 Flow-volume loops at baseline. Negative pressure-driven forced expiration maneuvers were performed at baseline to generate flow-volume(FV) loops. Mean FV loops are shown for each group. Bleo-fibrosis and PPE-emphysema, both parenchymal disorders, are shown in (a), LPS-ALI and
HDM-asthma are represented in (b), together with the naive control group. n = 7 –8 per groupDevos et al. Respiratory Research (2017) 18:123

aPage 8 of 14

Forced expiratory volume 0.1 s (FEV 0.1)

b

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P P leoE- -fibe m roph sis
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em
a

B

N

ai
ve

C

tr

ls

0

e 50

Peak expiratory flow (PEF)
*

***

40

mL/s

4

***
30
20

LI
H
D
M
-a
st
hm
a

LP
SA

P P leoE- -fibem roph sis
ys
em
a

B

N

ai
ve

C

tr

ls

10

Fig. 5 FE-derived parameters at baseline. Forced expiratory volume at 0.1 s, FEV0.1 (a), forced vital capacity, FVC (b), FEV0.1/FVC ratio (c), forced
expiratory flow at 0.1 s, FEF0.1 (d), and peak expiratory flow, PEF (e) were measured in mice by negative pressure-driven forced expiration. Parameters areshown for each individual mouse, along with group means. * p < 0.05, ** p < 0.01 and *** p < 0.001 compared to the naive control group. n = 7 –8 per group

trend in PEF and in FVC relative to control micewith increasing concentrations of MCh (Fig. 7a,c).
As for the associated NPFE-related parameters, allgroups showed a decreasing trend in FEV0.1 and/or
FVC with increasing concentrations of MCh (Fig. 8).
In mice with ALI, FEV0.1 was significantly reduced at
0, 0.6, 1.25 and 5 mg/mL of MCh, compared to control mice (Fig. 8a) although the differences between

the two groups were small. Also, FVC was significantly reduced over a similar range of MCh concentrations (Fig. 8b). The FEV0.1/FVC ratio of mice with
ALI did not differ from naive controls (see Additional
file 3: Figure S2). In mice with HDM-induced asthma,both FEV0.1 and FVC were significantly lower at all
MCh concentrations compared to control mice
(Fig. 8). The curves of FEV0.1/FVC ratio of mice

Devos et al. Respiratory Research (2017) 18:123

aPage 9 of 14b

Airway resistance LPS-ALI

1.2

Rn (cmH2O.s/mL)Rn (cmH2O.s/mL)

1.2
0.9
0.6
0.3

Airway resistance HDM-asthma
***

0.9

**
0.6
0.3
0.0

0.0
baseline 0 2.5 5.0

10.0

baseline 0 2.5 5.0

20.0

Naive Ctrls

10.0

20.0

Methacholine (mg/mL)

Methacholine (mg/mL)

Naive Ctrls

LPS-ALI

HDM-asthma

Fig. 6 Airway responsiveness to methacholine. Airway resistance (Rn) was measured in response to increasing aerosol challenges of methacholine(0 –20 mg/mL). Mean (± SD) concentration-response curves of Rn are shown for LPS-ALI in (a) and HDM-asthma in (b), and represented together
with that of the naive control group. ** p < 0.01 and *** p < 0.001 compared to the naive control group.n = 7 –8 per grouptreated with HDM, followed the curve of control
mice (see Additional file 3: Figure S2).
The provocative concentration of MCh required to induce a 20, 30 or 40% decrease in baseline FEV0.1, wassignificantly lower in mice with HDM-induced asthma
compared to naive controls (Fig. 9). In mice with ALI,
no changes in provocative concentrations of MCh were
found, compared to naive controls (data not shown).

ab

FV-loop Naive Ctrls
Baseline
0 mg/mL
0.625 mg/mL
1.25 mg/mL
2.5 mg/mL
5 mg/mL
10 mg/mL
20 mg/mL

40
30
20
10
0
0.0

0.5

1.0

Baseline
0 mg/mL
0.625 mg/mL
1.25 mg/mL
2.5 mg/mL
5 mg/mL
10 mg/mL
20 mg/mL

40
30
20
10
0
0.0

1.5

0.5

1.0

1.5

Volume (ml)Volume (ml)cFlow (mL/s)

Flow (mL/s)

FV-loop LPS-ALI
50

50

FV-loop HDM-asthma
50

Flow (mL/s)

Discussion
The lung carries many functions within the body, the mostimportant being that of providing a constant supply of O2 /removal of CO2 through the process of respiration. Thisfunction of the lung is essentially a mechanical one, as it relies on changes in pressure and volume. While it is highly
influenced by the mechanical properties of the respiratory
system, the lung can very often continue to carry it out

Baseline
0 mg/mL
0.625 mg/mL
1.25 mg/mL
2.5 mg/mL
5 mg/mL
10 mg/mL
20 mg/mL

40
30
20
10
0
0.0

0.5

1.0

1.5

Volume (ml)Fig. 7 Flow-volume loops. Negative pressure-driven forced expiration maneuvers were performed at baseline and following each methacholine
aerosol challenge (0-20 mg/mL) to generate 8 successive FV-loops for each group. Mean FV-loops are shown for Naive Ctrls in (a), LPS-ALI in (b)and HDM-asthma in (c).n = 7 –8 per groupDevos et al. Respiratory Research (2017) 18:123

aPage 10 of 14b

FEV0.1 LPS-ALI
1.5

1.5

**
*

mL

1.0

mL

FVC LPS-ALI
2.0

1.0

*

0.5

**
*** ** **

**

*

0.5

0.0

0.0

baseline 0 2.5 5.0

10.0

20.0

baseline 0 2.5 5.0

Methacholine (mg/mL)
Naive Ctrls

c

10.0

20.0

Methacholine (mg/mL)

LPS-ALI

Naive Ctrlsd

FEV0.1 HDM-asthma
1.5

LPS-ALIFVC HDM-asthma
2.0
1.5

0.5

****
**
***

mL

mL

1.0

1.0

***

0.5

***

******
***
***

***

***

0.0

***

***

0.0
baseline 0 2.5 5.0

10.0

20.0

baseline 0 2.5 5.0

10.0

20.0

Methacholine (mg/mL)

Methacholine (mg/mL)

Naive Ctrls

Naive Ctrls

HDM-asthma

HDM-asthma

Fig. 8 FE-derived parameters. Negative pressure-driven forced expiration maneuvers were performed at baseline and following each methacholine
aerosol challenge (0 –20 mg/mL). Forced expiratory volume at 0.1 s, FEV0.1 (a, c), forced vital capacity, FVC (b, d) are shown for LPS-ALI (a, b) and HDMasthma (c, d). The mean (± SD) concentration-response curve of each group is represented together with that of the naive control group. * p < 0.05, **p < 0.01 and *** p < 0.001 compared to naive controls. n = 7 –8 per group

under tidal volume conditions despite significant histologic
disease manifestations. Therefore, it is frequently necessary
to study the lung under different conditions in order to detect pathological lung function defects. In the present study,
we performed FE measurements in four well-establishedmouse models of respiratory diseases with the objective offurther assessing disease-specific changes using FE-derived
parameters. Studying the lung under forced expiration assesses its function under conditions susceptible of inducing
expiratory flow limitation. Combined with measurementsof respiratory mechanics, specific lung volumes, and quasistatic pressure-volume curves, FE measurements canpotentially add valuable information and contribute toproduce a more comprehensive functional assessment. Thisapproach, which explores the linear as well as the nonlinear behaviors of the lung, might be particularly important in mouse models, given the relatively large airways thatmice have for their body size [17, 18].
In the present study, all these measurements were
performed at baseline as well as in response to MChprovocations, in order to study the lungs under agreater level of functional impairment. The measurement device used offered a refined study design inthe sense that all measurement types could be madeseamlessly in the same subject, with no need to moveit during the course of the experimental protocol orother inconveniences associated to the use of separate
pieces of equipment [11]. In addition, the raw signals
generated during the forced expiration maneuver wereof improved quality relative to previously publishedwork [11].
FOT measurements at baseline proved to be useful toassess lung function in models of pulmonary fibrosis[13] and pulmonary emphysema [12, 14] and confirm
the expected changes in respiratory mechanics. The restrictive pathology, observed in mice with fibrosis, wasalso reflected in FE-related parameters. FEV0.1, FVC andPEF were significantly decreased in mice with fibrosis,
and could, thus, serve as diagnostic parameters havingsome similarities with those routinely used in the clinic.
This is in contrast to the earlier study of Vanoirbeek et al.,
where less changes in FE-related variables could beobserved in mice with fibrosis [11]. In line with the current
study, Tanaka et al. reported an increase in respiratory elastance and tissue elastance, as well as a decrease in FVC inmice with bleomycin-induced pulmonary fibrosis [19].

Devos et al. Respiratory Research (2017) 18:123

aPage 11 of 14b

PC20 calculation

100

80

PC20
70
60

mg/mL (Mch)

Naive Ctrls
HDM-asthma90

FEV0.1 (%)

PC20

6

50

*

4

2

0

0.0

2.5

5.0

10.0

20.0

Naive Ctrls

HDM-asthma

Methacholine (mg/mL)cdPC30 calculation

100

Naive Ctrls
HDM-asthma

80
70

PC30
60

mg/mL (Mch)90

FEV0.1 (%)

PC30
20

50

15
10

*
5
0

0.0

2.5

5.0

10.0

20.0

Naive Ctrls

HDM-asthma

Methacholine (mg/mL)ef

PC40 calculation

80
70
60

mg/mL (Mch)

Naive Ctrls
HDM-asthma90

FEV0.1 (%)

PC40
> 20

100

15

***

10
5

PC40
50

0
0.0

2.5

5.0

10.0

20.0

Naive Ctrls

HDM-asthma

Methacholine (mg/mL)

Fig. 9 Provocative methacholine concentrations in mice. Mean FEV0.1 concentration-response curves to MCh are shown for HDM-asthma and naive
controls, with indication of corresponding provocative concentration inducing a 20% (PC20) (a), 30% (PC30) (c), or 40% (PC40) (e) decrease in baseline
FEV0.1. PC20, PC30, and PC40, calculated from the concentration-response data for each individual mouse and group means are shown in b, d, and f,
respectively. * p < 0.05, *** p < 0.001 compared to naive control group. n = 7 –8 per group

Characterization of the pathology of pulmonary emphysema in mice treated with PPE showed the expected
decrease in G and H, and an upward shift in the PV
loop, reflecting decreased elastic recoil. Consistently,
there was an increase in the inspiratory capacity and adecrease in PEF, but no significant change in Cst in
our model, compared to naive healthy mice. Furthermore, in contrast to the human disease situation,
FVC was increased, resulting in a decreased FEV0.1/
FVC ratio [20, 21]. These results correspond withthose of Tanaka et al., who also assessed lungmechanics and respiratory dysfuntion in mice with
PPE-induced pulmonary emphysema, in terms oftissue elastance and FEV0.05, using FOT and NPFE
techniques, respectively [22]. Similar to our results,
they detected a pronounced decrease in both tissue
elastance and FEV0.05 under baseline conditions. The

lack of change in Cst observed in our study could be
related to the model used, the style of the PV loop
used (ramp versus plateau style), or alternatively, tothe fact that the PV loop was constructed over theinspiratory capacity range instead of over the full lungvolume range [23].
The FE-derived parameters discussed in this study,
although similar to those reported in humans, weregenerated by negative pressure. This methodology differsfrom the technique used with patients and could
possibly be responsible for some of the discrepancies
observed. In the PPE-treated mice, FEF0.1 was significantly increased compared to control mice and therefore
differs from what is typically observed in human subjects[20, 21]. Airway collapse and expiratory flow limitation
are important factors influencing forced expired flow
and these are sensitive to changes in the mechanicalDevos et al. Respiratory Research (2017) 18:123

properties of the respiratory system. As shown by a significant decrease in H, PPE-treated mice displayed anincreased lung compliance which, under negative
pressure-driven forced expiration, most likely led to thecreation of a choke point at a higher location in the airway
tree than in the control mice, thus resulting in a higher
upstream airway calibre and therefore a higher expiratory
flow at the selected time and negative pressure.
We also included a model of acute lung injury (ALI),
induced by endotoxin (LPS) exposure and characterized
by acute neutrophilic inflammation. FEV0.1 remained
unchanged in this mouse model. However, a significant
increase in Rn was observed in the LPS-ALI mice compared to control ones, as well as a significant decrease in
PEF, demonstrating the value of these measurements.
As shown before [11], mouse models of airway inflammation and airway hyperresponsiveness often display normal
values for respiratory mechanics parameters measured
under baseline conditions. Similarly, Rn and H were not affected in mice with HDM-induced asthma. Also in parallelto the study of Vanoirbeek et al. [11], Cst and IC did not
differ from naive controls under baseline conditions.
As expected, enhanced MCh airway responsiveness
relative to control mice could be demonstrated in micetreated with HDM (i.e. increased central airway resistance, Rn). In mice with ALI, the absence of MCh hyperresponsiveness in terms of Rn is in line with the findingsof Secher et al. [24] in male C57BL/6 mice. However,this latter study contradicts the work of Card et al. [25],in which it was reported that LPS-treated male C57BL/6
mice, and not female, exhibit an exaggerated MChevoked airway response. There are some small technicaldifferences that distinguish these studies, however, thelack of AHR in the present model of neutrophilic inflammation further confirms the existence of a dissociation between recruitment of inflammatory cells and thedevelopment of AHR [26–28].
In this study, we also measured airway responsivenessto MCh in terms of FE-derived variables. In contrast to
the study of Shalaby et al. [10], we did not perform adeep inflation after each FE measurement to reverse thederecruitment of lung units caused by the NPFE maneuverin order to be able to measure airway responsiveness in the
HDM-asthma model over a low methacholine concentration range. Mucus is often present in the airways of allergically inflamed mice and methacholine is a known mucus
secretagogue [29]. That technical difference most likely
enabled us to reach a greater level of airway functional
impairment at any given challenges by limiting the subsequent amount of aerosol delivered to the peripheral airways
and thus favoring a more proximal deposition. Despite ofthese differences in protocols, the shape of the FV loops
generated in the present study appeared generally similar to
those previously reported, with a typical MCh-evoked

Page 12 of 14

cumulative decrease in FEV0.1 and FVC. The FV loops
decreased gradually, with a more concave shape withincreasing MCh concentrations, reflecting an increasing
degree of airflow obstruction. The assessment of airway
responsiveness using the clinically-resembling parameters
FEV0.1 or FVC revealed functional changes in the HDMmodel of asthma studied, as well as in the LPS-ALI one.
Although in that latter model, the separation between
groups was much smaller compared to the HDM-model, as
also seen with the FOT technique. The difference in thelevel of statistical significance observed between the two
measurement techniques in the LPS-ALI model could be
related to the fact that small group sizes were analyzed or,alternatively, to the possibility that FOT and FE measurements are influenced by different determinants or determinant distributions [10]. In agreement with the latter
point, there was no statistical difference between the LPSALI and control groups when FE results were normalized
to the 0 mg/ml (saline) challenge (data not shown). The
FEV0.1 response curves also allowed, at least in the HDMinduced asthma model, the calculation of the provocative
MCh concentration inducing a 20 (PC20), 30 (PC30), or
40% (PC40) decrease in FEV and a classification of thesubjects’ AHR response as either mild, moderate, or severe,as performed clinically with human data [30, 31].
The addition of FE measurements to other types of lungfunction evaluations (e.g. FOT, PV loops, specific volumes)
contributed, in the present study, to produce a more
complete functional characterization of four well established respiratory disease models, all generated on the same
genetic background. This approach helped confirm disease
specific phenotypes while further assessing distinctive
changes using a number of clinically-resembling parameters. The translational utility of these latter parameters tohuman spirometry still remains to be established and theirinterpretation is also not fully understood. However, nonlinear behaviors of the lung, such as expiratory flow limitation, often play important roles in respiratory diseases andtheir evaluation should be, in our view, more frequently
included in the evaluation of preclinical models as well as
extended. For example, future studies could consider, morechronic models, different genetic backgrounds, as well as
additional lung function measurements (e.g. additional lungvolumes) or conditions under which measurements are
made (e.g. different PEEP or negative pressure).

Conclusion
In conclusion, four well established mouse models of
respiratory diseases were characterized with the objective of further assessing disease-specific changes using
FE-related parameters. The results showed that lungfunction defects could be detected in each disease modelby at least one FE-derived parameter under baseline or
bronchoconstricted conditions and that the changes

Devos et al. Respiratory Research (2017) 18:123

observed generally behaved in a similar manner as inhuman patients. Routinely exploring the linear and nonlinear lung behaviors in order to produce more comprehensive and insightful lung function assessments inmouse models could be key in increasing the translational utilities of preclinical studies.

Additional files
Additional file 1: Figure S1. Experimental design of different airway
disease models. Mice were sacrificed at the age of 13-14 weeks (indicatedin red). (PPTX 111 kb)
Additional file 2: Table S1. Complete overview of all lung function
parameters, measured at baseline using the flexivent FX. (DOCX 23 kb)
Additional file 3: Figure S2. Forced expiration-derived parameters.
Negative pressure-driven forced expiration maneuvers were performed atbaseline and following each methacholine aerosol challenge (0 –20 mg/mL). FEV0.1/FVC ratio is shown for LPS-ALI (a) and HDM-asthma (b). The
mean (± SD) concentration-response curve of each group is represented
together with the naive control group. n = 7 –8 per group. (PPTX 131 kb)

Acknowledgements
The project was supported by a grant from the Research Council of the KU
Leuven (PF-TRPLe) and by a grant of the Flemish Research Foundation(research grant 1.5.049.12 N).
Availability of data and materials
All data generated or analyzed during this study are included in thispublished article (and its supplementary information files)
Authors’ contribution
Conception and design – FD, JV, PH; Analysis – FD, AM, SS, LP, CAL, EV;
Interpretation – FD, AR, EV, BN, PH, JV; Drafting manuscript – FC, JV; Proof
reading – BN, AR, PH, RL, MT, SS, LP, AM. All authors read and approved thefinal manuscript.
Competing interests
The authors declare that they have no competing interests.
Consent for publication“Not applicable”
Ethics approval
All experimental procedures performed in mice were approved by the KU
Leuven local Ethical Committee for animal experiments (P166-2012)
Disclosure
Annette Robichaud is employed by SCIREQ Inc, a commercial entity involvedin a subject area related to the content of this publication. SCIREQ is anemka Technologies company.

Publisher’s NoteSpringer Nature remains neutral with regard to jurisdictional claims inpublished maps and institutional affiliations.
Author details
1
Center for Environment and Health, KU Leuven, Leuven, Belgium. 2Molecular
and Medical Virology, Ruhr-University, Bochum, Germany. 3SCIREQ Scientific
Respiratory Equipment Inc., Montreal, QC, Canada. 4Clinical Immunology, KU
Leuven, Leuven, Belgium. 5Skeletal Biology and Engineering Research Center,
KU Leuven, Leuven, Belgium. 6Translational Cell and Tissue Research, KU
Leuven, Leuven, Belgium. 7Institute of Clinical and Molecular Virology,
University Hospital Erlangen, Erlangen-Nürnberg, Germany.

Page 13 of 14

Received: 13 February 2017 Accepted: 12 June 2017

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