2. M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171 S153
great progress from these models, their application is limited
by the considerable practical and financial issues involved.
This paper will focus on CF mouse models, presenting an
update on new developments and novel data, in particular on
the properties of the mutant strains made available to the CF
community through the EuroCareCF project.
2. CF mutant mice, portrait of an unruly family
Shortly after the discovery of the CFTR gene, several
groups set out to generate mouse strains with mutations in the
Cftr locus using the homologous recombination technology
and mouse embryonic stem cells pioneered by Mario Capec-
chi, Martin Evans and Oliver Smithies who jointly received
the Nobel Prize for these innovations in 2007. Table 1 lists
the genetically modified mouse strains produced by different
groups. Details of mouse strain nomenclature and availability
can be found at the Mouse Genome Informatics website
(http://www.informatics.jax.org/). Table 2 summarizes briefly
the major phenotypic traits of CF mice in comparison with the
human disease.
Several loss of function, or “knock-out” models were made
by interrupting the mouse Cftr coding region with a selectable
marker. The Cftrtm1Hgu shows low level of residual activity
of wild-type CFTR and lower mortality, compared to the
other two frequently used knockout strains Cftrtm1Cam and
Cftrtm1Unc (Table 1).
In addition, common CF causing mutations were intro-
duced into the mouse CFTR amino acid sequence. Three
strains with the F508del mutation in the mouse Cftr gene
were generated, two of which carry an intronic insertion that
may interfere with gene transcription (Table 1). By contrast,
the Cftrtm1Eur allele was made by the two step “hit and
run” recombination procedure, which removes the selectable
marker. Therefore, this allele only contains the F508del mu-
tation and has normal transcription rates in all tissues [9,10].
All three strains have been successfully used in phenotypic
studies. The F508del mutation in the mouse Cftr context
causes severely reduced amounts of fully glycosylated CFTR
protein (band C) similar to the human form, and a phenotype
comparable to CFTR loss of function mutants [9]. In cultured
gallbladder epithelial cells from F508del mutant mice, the
number of active CFTR Cl− channels observed in patch-
clamp experiments is increased by incubating cells at low
temperature [9], suggesting a similar temperature-dependent
processing defect to the F508del mutation in human CFTR.
Moreover, the G551D mutation which disrupts channel gating
has the typical CF phenotype caused by loss of function
mutants [11]. The less severe phenotype of the F508del-
and G551D-CFTR models when compared to CFTR knock-
outs, particularly lower perinatal mortality due to intestinal
obstruction, has been attributed to residual CFTR activity.
To overcome the limitations imposed by the severe in-
testinal disease presented by many knockout strains, “gut
corrected” hybrid strains were created using a transgene ex-
pressing CFTR from a promoter specific for the intestinal
epithelium (Table 1).
The most recent and important addition to the CF mouse
family is a conditional (“loxed”) Cftr deletion that can be
created by cell specific expression of the cre recombinase [12].
This genetically modified mouse will allow the contribution
of different cell lineages to CF pathology to be elucidated.
Although not a CFTR mouse model, genetically modi-
fied mice overexpressing the β subunit of the epithelial Na+
channel (ENaC) are important because they possess a lung
phenotype resembling that of CF [13–16]. For further infor-
mation about this mouse model and its applications, see Zhou
et al. [17].
3. Mouse models, sources of variation and practical
considerations
CF mutant mice are generally much less fertile than normal
mice, which makes it difficult to breed them from homozy-
gous mutant pairs. This is partly due to frequent obstruction
of the male vas deferens [18,19]. However, homozygous fe-
males are also poor breeders. Occasionally mice can be bred
successfully from homozygous pairs. However, there is risk
of “genetic drift”, the selection of compensating mutations in
either the background or the Cftr allele, which affect pheno-
type in an unpredictable way. Breeding CF mutant mice from
heterozygous animals is therefore recommended strongly. Im-
portantly, this breeding strategy yields age- and sex-matched
normal littermates, which are mandatory control for most
experiments. Breeding requires considerable investment in
DNA genotyping of all neonates by ear or tail clips, the
identification of individual animals by chip, toe- or ear-clips
and a database to track individual animals. Genotyping is
primarily required to identify heterozygous breeders, because
homozygous mutants are easily recognized by their typical
white incisor teeth [20].
There is consensus about the most prominent phenotypic
abnormalities of different Cftr mutant mice. Yet, close com-
parison of published data from different laboratories shows
considerable variation and even controversy about the nature
and extent of lung disease. One obvious source of variation
is the different Cftr mutations, some of which are associated
with residual CFTR function (Table 1). Furthermore, Cftr
alleles have been bred onto different genetic backgrounds,
which may cause subtle differences in phenotype, presumably
because of the expression of different “modifier genes”. Com-
plete genomes of several laboratory mouse strains are now
available (for further information, see Sanger mouse genome
project, http://www.sanger.ac.uk/resources/mouse/genomes/).
This resource will provide a new and powerful tool to ana-
lyze phenotypic differences between CFTR mutant strains. It
will also facilitate comparative studies of modifier genes and
potential therapeutic targets in the human population.
Animal husbandry is a critical issue, especially when in-
vestigating innate adaptive responses (e.g. lung inflammation),
which are by their very nature designed to reflect external
conditions. This can cause differences between strains, con-
tribute to inter-individual variation, and lead to differences
in results between laboratories. Unfortunately, there is no
3. S154 M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171
Table 1
CFTR mutant mice
Mouse Mutation Cftr mRNA Genetic Survival to Body wt Contact References
background maturity
Null mutations
Cftrtm1Unc S489X Ex10 R Not detectable* C57Bl/6 <5% 10–25% reduction BH Koller/Jax Labs [113,158]
Cftrtm1Cam R487X Ex10 R Not detectable 129S6/Sv/Ev <5% 20% reduction WH Colledge [159]
Cftrtm1Hsc M1X Ex1 R Not detectable CD1 x 129 25% Delayed LC Tsui [160]
Cftrtm3Bay Ex2 R Not detectable C57Bl/6 x 129 40% Reduced AT Beaudet [161]
Cftrtm3Uth Y122X Ex4 R Not detectable C57Bl/6 25% 25–50% reduction M Capecchi/PB Davis [113,162]
Hypomorphic mutations
Cftrtm1Hgu ** Ex10 I 10% of wt MF1 x 129 90% No reduction J Dorin [113]
Cftrtm1Bay Ex3 I <2% wt C57Bl/6 x 129 40% 70% reduction AL Beaudet [163]
F508del mutations
Cftrtm2Cam F508del R 30% of wt 129S6/Sv/Ev <5% 10–20% reduction WH Colledge [164]
Cftrtm1Kth F508del R Low in intestine C57Bl/6 x 129 40% 10–50% reduction KR Thomas/Jax labs [165]
Cftrtm1Eur F508del (H&R) Normal levels FVB/129; FVB 90% 10–20% reduction BJ Scholte [9]
C57Bl/6
Other point mutations
Cftrtm2Hgu G551D R 50% of wt CD1/129 65% 30–50% reduction J Dorin [11]
Cftrtm3Hgu G480C (H&R) Normal levels C57Bl/6/129 Normal No reduction J Dorin [166]
Cftrtm2Uth R117H R 5–20% of wt C57/Bl6 95% 10–25% reduction M Capecchi/PB Davis [113,162]
Transgenes
Mouse Transgene Promoter Expression Phenotype References
Tg(FABPCFTR) CFTR Rat intestinal fatty acid Intestinal villus epithelia Rescue of CF intestinal pathology [167]
binding protein
Tg(CCSPScnn1b) Scnn1b Clara cell secretory Airway surface epithelia Na+ hyperabsorption [13]
protein (CCSP) Reduced airway surface fluid volume
Mucus accumulation, CF-like lung disease;
40% survival.
Legend: Mutations are introduced by homologous recombination in embryo stem cells. Key: R, replacement; I, insertion; (H&R), Hit & Run; * a splice variant
was detected [168]. Transgenes are created by injection into mouse oocytes. General rules on mouse gene and strain nomenclature can be found at the Mouse
Genome Informatics guidelines website.
** The strain Cftrtm1Hgu is also referred to as CftrTgH(neoim)Hgu due to the unique nature of this recombinant which produces low level normal CFTR due to a
splice variant introduced by the recombination event.
global standard protocol for animal husbandry. Laboratories
therefore differ in the level of pathogen exposure. The Rot-
terdam animal facility maintains a FELASA+ pathogen-free
standard using sterilized ventilated cages supplied with steril-
ized, acidified water. Separation of animals causes stress and
is avoided whenever possible; nesting material (e.g. sterile
paper tissue) is always provided. Strict light and dark regimes
are maintained; nursing females are not disturbed until ten
days after birth to reduce postnatal mortality. CF homozygous
mutant animals do not travel very well and often die or
suffer excessive weight loss during long distance transport.
Therefore, animals are moved at least two weeks in advance
of experiments to allow them to adapt. Under these condi-
tions we can maintain colonies of Cftrtm1Eur F508del and
Cftrtm1Cam knockout mice in different genetic backgrounds
on normal chow at less than 20% mortality for homozygous
mutants.
Gender is another source of variation that should not be
underestimated. Male and female mice differ with respect to
many physiological responses relevant to CF studies. This
is well illustrated in a recent study of a mouse model of
asthma [21]. Gender differences should be taken into account
when designing experiments and analyzing data. Finally, age
is another important variable. Its importance is demonstrated
by the progressive nature of lung disease in several CF mouse
strains. Together, these considerations place severe demands
on the size and selection of experimental groups necessary to
generate reproducible data and statistical power.
4. CF mouse phenotype
The major phenotypic abnormalities of CF mutant mice
are briefly summarized in Table 2. This brief overview does
not do justice to all the available literature.
In the mutant strains investigated, pancreatic pathology
is not a prominent feature, although it was reported in a
C57Bl/6 Cftr knockout backcross [19]. Although pancreatic
insufficiency correlates with severe pathology, it is not seen
in all CF patients. Similarly, male infertility caused by
obstruction or absence of the vas deferens is frequent in
CF mice depending on the strain and genetic background
[18,19]. However, this pathology appears less severe than in
humans where absence of the vas deferens is observed in
all male CF patients. Apparently, factors other than CFTR
4. M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171 S155
Table 2
Comparative CF pathology in human and mouse CF mutants
Human pathology [27,169] Tissues affected in
CFTR mutant
Mouse pathology [1,2]
Recurrent bacterial lung infections,
chronic inflammation, airway
remodeling, irreversible loss of
function
[170] Lung No CF type obstructive lung disease.
Inflammation and tissue remodeling in aging
animals.
Hyper-inflammation reduced clearance upon
challenge with bacteria or LPS.
Pro: [19,105,114,
117,171,172]
Con: [173]
Hyposecretion, viscous mucus,
plugging
[174,175] Submucosal gland
(nasal, bronchial)
Nasal and tracheal glands present, reduced
cAMP-induced secretion in bronchial glands of
mutant mice
[81,82]
Bacterial biofilm, viscous/purulent
mucus, defective mucociliary
clearance, goblet cell hyperplasia,
inflammation, tissue remodeling
[176,177] Trachea, bronchial
epithelium (ciliated/
goblet)
Reduced mucociliary clearance (contested).
Electrophysiology influenced by CaCC
(TMEM16A).
Pro [84]
Con: [83]
Distal airways involved in early
disease, CFTR expression in Clara
cells
[95] Broncheoli (distal
epithelium, Clara cells)
Spontaneous progressive remodeling and
inflammation (strain- and age-dependent)
[19,102,172,178]
No known pathology, CFTR
expression in cultured type II cells
Alveoli (type I/II) Cftr expression in type II cells, abnormal fluid
secretion in mutant mice.
[97,99].
Meconium ileus, chronic
malabsorption, reduced body
weight, failure to thrive (pancreas
deficiency), typical
electrophysiology.
[31,179,
180]
Intestine Lethal mucus plugging, lipid and bile acid
reabsorption defect, reduced body weight, failure to
thrive. Goblet cell hyperplasia in ileum, mucus
accumulation in intestinal crypts, meconium ileus
equivalent, chronic inflammation, abnormal
electrophyiology.
[1,2,25,60,181,
182]
Frequent biliary disease,
progressive cirrhosis
[43,183–
185]
Liver/Biliary duct No morbidity, progressive liver disease in aging CF
KO mice biliary duct damage in KO mice upon
induced colitis
[19,186,187]
No frequent pathology [188] Kidney Functional CFTR in proximal tubule. In CF mice no
overt pathology, abnormalities in Na+ and K+
transport, defective endocytosis.
[41,42,189,190]
Increased frequency of bile stones Gallbladder No cAMP-induced Cl− or water secretion [45,191]
Pancreatitis, atrophy, digestive
enzyme secretion deficiency,
diabetes, abnormal bicarbonate
secretion and mucous plugging of
ducts
[192,193] Pancreas Normal function and ductal fluid secretion,
occasional duct plugging in Cftrtm1Unc mice
[19,194]
No pathology Tooth Abnormal enamel (white incisor) [20]
Osteopenia, osteoporosis [73,74] Bone Abnormal bone density. [67–69]
No pathology, reduced flow rate [195] Salivary gland Adrenergic fluid secretion defective [38]
High NaCl in sweat (defective
reabsorption duct)
[169] Sweat gland Foot pad glands only, no report of deficiency. [196]
Absent (CBAVD, male infertility
100%)
Vas deferens Frequent plugging or absent (male infertility) [18,19]
Frequent obstruction Oviduct Frequent obstruction (female infertility)
No pathology reported Cornea Defective chloride permeability, enhanced amiloride
response, attenuated uptake of Pseudomonas
[197–199]
Smooth muscle cell-related
pathology in lung and diaphragm
suspected but not firmly established
Smooth muscle cell CFTR expression, altered function and innervation,
degeneration in CF KO mice
[47–50,200]
No pathology reported Mast cells CFTR expression, enhanced presence in CF KO
intestine, no evidence for cell autonomous
abnormality
[59,60]
No pathology reported Macrophages CFTR expression, reduced capacity to kill
Pseudomonas, abnormal proinflammatory cytokine
secretion
[51,55–57]
5. S156 M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171
deficiency determine disease severity, in this case probably
the expression of alternative pathways for Cl− transport.
4.1. Intestinal disease in CF mouse models
Similar to humans, intestinal disease is prominent in all
CF mutant mice. Depending on the strain and laboratory
conditions many mutant mice may die of intestinal blockage
around the time of weaning. This is a stochastic effect at-
tributed to abnormal fluid and mucus secretion (meconium
ileus equivalent). To reduce the frequency of intestinal ob-
struction, mutant mice are often fed a liquid diet (Peptamen™)
[22]. Because this drastic dietary intervention may affect the
phenotype of both control and mutant mice, the use of a
mild laxative (Poly Ethylene Glycol; PEG) in combination
with normal chow is recommended as an alternative [23].
In the Rotterdam facility, Cftrtm1Eur and Cftrtm1Cam strains
(FVB/n and C57Bl/6) survive well into adulthood (10–30
weeks, 80–90%) on normal chow based on animal fat (Hope
Farm™ SRMA 2131, Abfarm, Woerden, the Netherlands)
combined with 6% (w/v) PEG in the drinking water for the
knockout mice. Male and female mutant mice of all strains
show a mild, but significant (10% average), reduction of body
weight compared to normal age- and sex-matched littermates.
Although there is no evidence of an essential lipid deficiency
[24], mutant mice display reduced lipid reabsorption, which is
most pronounced in the Cftrtm1Cam strain [25]. Interestingly,
the length and weight of the small intestine is larger in mutant
Cftrtm1Eur C57Bl/6 compared to wild-type (10%, N = 18,
P < 0.01, Wilke et al., unpublished data [26]).
In the mouse and human intestine, different proximal
to distal regions with distinct morphological and functional
characteristics can be distinguished. In the duodenum, bile
from the gallbladder and digestive enzyme secretions from the
pancreas are delivered through a common duct and mixed with
the acidic products of the stomach. In the jejunum and ileum,
regulated fluid and mucus secretion maintain the optimal
milieu required for lipid, sugar and amino-acid absorption.
The colon, including the caecum, is involved in mucus and
bicarbonate secretion, and net ion and fluid reabsorption,
compacting the intestinal content to faeces. In addition to
proximal to distal gradients along the length of the intestine,
the epithelium is organized vertically by invaginations called
crypts and finger-like protrusions called villi. Stem cells at the
bottom of the crypts continuously produce new epithelial cells
that differentiate to form the different cell types, lining crypts
and villi.
CFTR is expressed mainly in enterocytes where it is
involved in chloride and bicarbonate secretion. The role of
CFTR in intestinal ion transport [27,28] and in the small
intestine [29,30] has been reviewed. The ion transport prop-
erties of the colon, including the role of CFTR were reviewed
by Kunzelmann and Mall [31]. Here, we present general and
novel data relevant to this review on CF mouse models.
Figure 1 demonstrates that the jejunum, ileum and colon
of backcrossed Cftrtm1Eur C57Bl/6 and FVB/n mice exhibit
a normal structure of crypts and villi. In the ileum a
distinct hyperplasia of goblet cells is observed, comparable to
published data for the original outbred strain (FVB/129) [10].
Consistent with this observation, the expression of the goblet
cell marker Clca3/Gob5 is increased 3-fold in the intestine of
CF mutant mice [32] and individual goblet cells appear larger
than normal, possessing “plumes” of mucous material. Mucus
plugs are often seen in the crypts and lumen.
A recent study in Cftrtm1Kth F508del (C57Bl/6) mice
highlights the crucial role of CFTR dependent bicarbonate
secretion in PGE2-induced intestinal mucus secretion and hy-
dration [33,34]. In the proposed model, bicarbonate secretion
by CFTR in enterocytes is required for the proper expansion
of tightly packed mucus-calcium ion complexes, which are
secreted from vesicular granules by neighbouring goblet cells.
In the absence of sufficient CFTR activity, the mucus does not
unfold properly, which might play an important role in CF
pathology. This indirect model for intestinal mucus secretion
was invoked because CFTR expression in goblet cells has
not been demonstrated directly, whereas enterocytes show
relatively high levels of CFTR expression. If it can be shown
that CFTR-mediated bicarbonate secretion plays a similar
role in mucus release in human airway epithelia, this would
have important implications for our understanding of CF lung
disease.
The mechanism causing the apparent goblet cell hyperpla-
sia in the ileum of CF mutant mice has not been established.
However, abnormalities of mucus composition in Cftrtm1Unc
knockout mice [35], suggest an effect on goblet cell differen-
tiation and function, possibly by an adaptive process.
Immunohistochemistry using the R3195 anti-CFTR anti-
body (CR Marino, University of Tennessee) shows CFTR
expression on the apical membrane of enterocytes in villi
and crypts of wild-type C57Bl/6 mice (Fig. 2). This signal
is strongly reduced in Cftrtm1Eur F508del mutant mice and
absent in Cftrtm1Cam knockout mice. Similar results were
reported using the original Cftrtm1Eur 129/FVB strain [9]
and the Cftr partial knockout Cftrtm1Hgu (also designated
CftrTgh(neoim)Hgu) [36]. Consistent with these data, electro-
physiological analyses demonstrate the absence or severe
reduction of CFTR function in CF mutant mice. For exam-
ple, Fig. 3 shows that in three different genetic backcrosses
of the Cftrtm1Eur F508del allele forskolin-induced (cAMP-
dependent) short-circuit current (Isc), a measure of CFTR-
mediated transepithelial ion transport, is reduced by more
than 80%. Figure 3 also reveals that the carbachol response
(Ca2+-dependent) is reduced, which in ileal epithelia reflects
the activity of Ca2+-activated K+ channels in the basolateral
membrane that facilitate CFTR-mediated Cl− secretion. In
CFTR knockout mice of the same genetic background, both
the forskolin and carbachol responses are completely lacking,
suggesting that the residual Isc response in intestinal epithelial
of Cftrtm1Eur F508del mice reflects residual CFTR activity
(De Jonge & Bot, unpublished).
As for human CFTR, the intracellular transport defect
and severely reduced activity of F508del-CFTR in Cftrtm1Eur
homozygous mice is rescued by low temperature incubation
[9]. Full restoration of wild-type CFTR protein processing
6. M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171 S157
Fig. 1. Goblet cells in CF mutant mouse intestine. Comparison of representative paraffin sections from mouse jejunum, ileum and colon stained with Alcian
blue to identify goblet cells. Wild-type C57Bl/6 (A) and FVB (C) are compared with their Cftrtm1Eur C57Bl/6 F508del (B) and Cftrtm1Eur FVB F508del (D)
homozygous mutant littermates (12–16 weeks). Ileum derived from both mutant strains (B and D) show significant hypertrophy of goblet cells. No differences
in jejunum and colon between wild-type and mutant mice are observed (original magnification 20×).
and channel activity was observed in stripped ileal mucosa
from Cftrtm1Eur homozygous mice after 12 hours incubation
at 26°C [37] (Wilke, de Jonge, in preparation). Based on
this observation, small molecules that rescue the cell surface
expression of mutant CFTR (termed CFTR correctors) are
now being tested in Cftrtm1Eur homozygous mice.
4.2. Salivary glands
Best and Quinton [38] first demonstrated that β-adrenergic
(cAMP)-stimulated fluid secretion by salivary glands is a non-
invasive and highly discriminative assay of CFTR function.
In this assay, atropine is first injected to prevent the stimula-
tion of secretion by cholinergic agonists before isoproterenol
is injected to activate β-adrenergic-stimulated fluid secretion.
Subsequently, Noël et al. [39] demonstrated that this assay can
be used to evaluate the efficacy with which small molecules
restore function to Cftrtm1Eur F508del mutants in vivo. This
assay offers the advantage of detecting very low levels of
residual function in mutant mice (Fig. 4). Moreover, it can be
performed repetitively in the same mouse, serving as a highly
sensitive indicator of CFTR function [39]. Other aspects of
the physiology of the salivary acini and glands in normal and
CF mutant mice were described recently by Catalan et al.
[40]. The authors’ data demonstrate that submandibular duct
cells co-express CFTR and ENaC and that CFTR is required
for effective reabsorption of NaCl [40].
4.3. Kidney
CFTR is expressed in the proximal tubules of the mouse
kidney, localized to the apical and endosomal compartments
of epithelial cells. Though no overt kidney pathology is
present, abnormalities in urinary protein secretion were ob-
7. S158 M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171
Fig. 2. Immunohistological analysis of CFTR expression in the ileum. CFTR protein was stained with the R3195 anti-Cftr antibody on paraffin sections.
Intense CFTR staining (brown) is observed in the ileum of FVB/n wild-type mice at the apical border of crypt and villus epithelial cells (A). Staining in
Cftrtm1Eur F508del FVB mutant littermates is much less intense and more diffuse (B). Tissue of the Cftrtm1Cam FVB knockout mouse serves as negative
control (C). This figure is representative for all backcrossed strains described in the present paper (original magnification 20×).
Fig. 3. Short-circuit measurements (Isc) of muscle-stripped excised ileum. Muscle stripped mucosa from ileum was analysed in an Ussing chamber as
described in [77]. Bars represent average initial short-circuit current (Isc) or the change of Isc after sequential addition of the cAMP agonist forskolin (forsk),
the Ca2+ agonist carbachol and glucose. Isc measurements were performed on ileum derived from the Cftrtm1Eur F508del strain in three different congenic
backgrounds (F13) A: FVB/n (F13) B: C57Bl/6 (F13) C: 129/sv (F12) comparing homozygous mutant (dd, open bars) with homozygous normal (++, black
bars). In all homozygous mutant Cftrtm1Eur F508del mice, forskolin responses are significantly lower than normal (P < 0.001), but residual activity is detected.
Similar results are observed with carbachol. Glucose is added to test a CFTR-independent function and to evaluate tissue viability. All animals were adult
males and females in the range of 13–30 weeks. Data presented are means ± SEM (n = number of mice, two pieces of ileum per mouse, FVB +/+, n = 9;
FVB dd, n = 10; C57Bl/6 +/+, n = 10, C57Bl/6 d/d, n = 15; 129sv +/+, n = 11, 129sv d/d, n = 8).
served in Cftrtm1Cam knockout mice, suggesting a role for
CFTR in membrane trafficking and brush border matura-
tion. To a lesser extent these abnormalities were also seen
in Cftrtm2Cam and Cftrtm1Eur F508del mice, where they are
attributed to partial processing of the mutant protein in this
tissue [41,42].
4.4. Liver
CF associated liver disease (CFLD) is an increasing
clinical problem, affecting more than 50% of adult CF
patients. CFTR is expressed in cholangiocytes, where it is
presumably involved in fluid homeostasis in the biliary duct
system [43]. In CF mice, no obvious liver pathology is
detected. Chronic administration of cholate increases weakly
bile production in CF mice when compared with control mice.
Moreover, by lowering the phospholipid-to-bile salt ratio,
chronic cholate administration increases the cytotoxicity of
bile produced by CF mice. By contrast, either acute or chronic
ursodeoxycholic acid (UDCA) administration induces a bile
salt-specific choleresis in vivo in mice that is independent
of functional CFTR [44]. These observations may help to
explain the beneficial effects of oral UDCA therapy in
cholangiopathies including CFLD.
4.5. The gallbladder
In mice, the gallbladder is an accessible and relatively
simple epithelial tissue with both secretory and absorptive
properties. We have previously shown that CFTR-dependent
forskolin-induced Isc ( Isc forsk) is absent in Cftrtm1Cam
C57Bl/6/129 knockout mice, but not normal mice [45,46]. We
8. M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171 S159
Fig. 4. Isoproterenol-stimulated salivary gland secretion. Salivary gland
secretion was measured in anesthetized mice for 21 minutes as described
in [38]. Mice were pretreated with atropine to block cross-stimulation of
cholinergic-stimulated secretion. Isoproterenol was used in the presence
of atropine to stimulate adrenergic secretion. There was no significant
difference between the maximum secretion of FVB wild-type (WT) male (n
= 6) and female mice (n = 4). The CF mutant homozygous Cftrtm1Eur FVB
F508del (DF/DF) littermates (DF/DF: male: n = 123, female: n = 63) and
homozygous Cftrtm1Cam FVB knockout mice (−/−: male: n = 7, female: n =
11) of either sex had a very low response compare to wild-type (P < 0.001).
The Cftrtm1Eur F508del mice do not differ from the Cftrtm1Cam.
further showed that unstimulated gallbladders from normal
mice reabsorb fluid to concentrate bile, whereas they secrete
fluid when stimulated with forskolin [45]. By contrast, in
gallbladders from Cftrtm1Cam knockout mice forskolin abol-
ished fluid reabsorption without stimulating fluid secretion,
demonstrating that fluid secretion is CFTR-dependent [45].
In Cftrtm1Eur 129/FVB F508del mice, forskolin-elicited Isc
responses were reduced by 80% [9]. Because Cftrtm1Cam
FVB mice also show low, but detectable, forskolin-elicited Isc
responses through a CFTR-independent pathway, the residual
forskolin-induced Isc is not mediated by F508del-CFTR (De
Jonge, Bot unpublished data). Here, we present previously
unpublished data demonstrating that the forskolin-induced
Fig. 5. Fluid transport in mouse gallbladder. Net fluid transport rates (Jv)
in isolated gallbladders of the 129/FVB Cftrtm1Eur F508del strain [10] were
measured as described in [45]. Negative values show initial transport of fluid
from the lumen of the bladder to the bath. After addition of the cAMP
agonist forskolin, Jv changes to positive values in normal mice, i.e. net
fluid secretion towards the lumen. By contrast, in homozygous Cftrtm1Eur
F508del littermates transport towards the bath is inhibited and there is no net
secretion into the lumen.
inhibition of reabsorption and absence of fluid secretion is
also observed in Cftrtm1Eur 129/FVB F508del mice (Fig. 5).
This indicates that CFTR activity is insufficient for net fluid
secretion in this CF mutant mouse. Gallbladder epithelial
cells contain a large number of secretory granules containing
mucins. Peters et al. [46] demonstrated that basal and ATP-
stimulated mucus release from cultured gallbladder epithelial
cells of Cftrtm1Eur 129/FVB mice are quantitatively normal.
However, the potential role of CFTR-mediated bicarbonate se-
cretion in mucus release in this model system and the physical
properties of the secreted mucus remain to be investigated.
4.6. Smooth muscle cells and CFTR function
CFTR is expressed in smooth muscle cells and skeletal
muscle myotubes, colocalised with the sarcoplasmic reticu-
lum [47–50]. Recently Divangahi et al. [47] demonstrated that
CFTR deficient myotubes display increased Ca2+ signaling
upon depolarisation and excessive NF-κB translocation in re-
sponse to inflammatory stimuli. Furthermore, degeneration of
diaphragmatic muscle was observed selectively in Cftrtm1Unc
mice challenged with Pseudomonas aeruginosa, which chron-
ically infects the lungs of CF patients [47]. The authors
suggest therefore that reduced muscle strength, particularly
of the diaphragm, as frequently observed in CF patients,
may contribute to CF lung disease. Because smooth muscle
cells of the airways and arteries express functional CFTR
[48–50], elements of the CF lung phenotype (e.g. wheezing,
hyperreactivity and progressive tissue remodeling) might also
result from the dysfunction of smooth muscle.
4.7. The cellular immune system is abnormal in CF mutant
mice
Several authors have reported abnormal behaviour of
alveolar and peritoneal macrophages in CF mutant mice. Di et
al. [51] demonstrated defective killing of bacteria after uptake,
possibly due to dysfunctional CFTR-dependent acidification
of endosomes/phagosomes [52], though this explanation is
contested [53,54]. Leal and co-authors showed that peritoneal
and alveolar macrophages from Cftrtm1Eur mice display a pro-
inflammatory bias after stimulation in culture, characterized
by enhanced secretion of IL1β, but reduced secretion of IL10
[55,56]. Similarly, Bruscia et al. [57] reported abnormal pro-
inflammatory cytokine secretion by lung and bone marrow-
derived macrophages in CF mutant mice after exposure to
lipopolysaccharide from Pseudomonas aeruginosa (PA LPS).
These data strongly suggest that macrophages from CFTR
mutant mice display defects that might, at least in part,
explain the pro-inflammatory lung phenotype in CF mutant
mice. Other cells of the myeloid lineage (e.g. neutrophils
[58]; mast cells [59,60] and dendritic cells [61]) are now
under investigation in CF patients and mouse models. If it
can be demonstrated that such defects are prominent in CF
patients and cell-autonomous, not adaptive, in origin, this
would support the conclusion that CF is a disease of the
immune system.
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4.8. Abnormal bone and cartilage formation in CF mutant
mice
Bonvin et al. [62] first showed that rings of tracheal carti-
lage in CF mutant mice (Cftrtm1Unc knockout and Cftrtm1Eur
F508del (129/FVB)) are frequently incomplete at birth. Be-
cause, neonatal CFTR knockout ferrets [6] and pigs [4,5] also
exhibit abnormal tracheal cartilage, this developmental abnor-
mality in CFTR deficient animals is the first to be verified in
multiple species. An explanation for this phenomenon is not
readily available. Interestingly, mouse chondrocytes, express
functional CFTR [63], suggesting a cell autonomous defect
in cartilage formation. However, a response to local and sys-
temic inflammation cannot be excluded. Of note, a similar, but
equally unexplained, phenotype was reported in TMEM16A
mutant mice [64–66], which lack the Ca2+-activated Cl−
channel (see above, section 4.12).
Reduced bone density in CFTR mutant mice was described
initially by Dif et al. [67] and confirmed by others [68,69].
It is currently unknown whether this defect is caused by
a cell autonomous CFTR-related defect of bone forming
osteoblasts or bone degrading osteoclasts. Alternatively, this
defect might be secondary to systemic inflammation and
abnormal bioactive lipid metabolism (ceramides, vitamin D).
Low amounts of CFTR protein are expressed in mouse
and human osteoblasts and odontoblasts, together with low
levels of CFTR mRNA [70]. The location of CFTR in these
cells was mainly intracellular, suggesting that CFTR might
function as an anion transporter in intracellular organelles.
The dysfunction of the ClC transporter ClC-7 in osteopetrosis
[71,72] raises the possibility that CFTR dysfunction in
organelle membranes of bone cells might underlie the changes
in bone metabolism in CFTR-deficient mice and CF patients,
leading to osteopenia.
The clinical relevance of these data remains to be estab-
lished. To date, no cartilage abnormalities have been reported
in CF patients. However, osteopenia and osteoporosis are
serious problems in CF patients [73,74]. Taken together, the
data suggest that animal model studies of CF-related bone
disease might lead to novel treatments for CF patients.
4.9. Nasal airways of CF mutant mice
The nasal epithelium is an important focus of CF mouse
model research, because of its similarities in structure and
function with the human bronchial airway epithelium. Indeed,
early studies of the nasal epithelium demonstrated electro-
physiological abnormalities consistent with CFTR dysfunc-
tion in CF mutant mice (i.e. abrogated cAMP-stimulated Cl−
conductance and enhanced amiloride-sensitive Na+ currents
in microperfusion studies, Table 2).
More recent studies draw attention to the complex ar-
chitecture of the mouse nasal epithelium and the effects of
CFTR dysfunction on this tissue [75,76]. The mouse nasal
cavity is a septated cartilage and bone structure covered with
nasal airway epithelium (NAE), mainly ciliated and mucus
producing (goblet) cells. However, the cavity lining the up-
per part of the skull is covered with olfactory epithelium
(OE). The OE consists of a multilayer of neuronal cells
projecting sensory cilia from the surface, covered with a
layer of specialized epithelial cells called sustentacular cells
(Fig. 6A and B). This tissue expresses CFTR and ENaC and
is thought to contribute to fluid and ion homeostasis across
the OE [76]. Under the OE and the NAE, there are numerous
submucosal glands embedded in connective tissue. CF mutant
mice display an age-dependent progressive degeneration of
the olfactory neuronal layer that is evident in adult mutant
mice from the age of 16–24 weeks. This was first reported
for Cftrtm1Unc knockout and CftrTgHm1 G551D mutant mice
on different genetic backgrounds [75,76]. Furthermore, an
increase of NAE thickness and apparent hyperplasia of mucus
producing cells is observed in CftrTgHm1 G551D mutants
[75]. At age 14–20 weeks, Cftrtm1Eur FVB mice did not show
histological evidence of OE degeneration in our facility (Fig.
6A, B), indicating that this abnormality is not only age-, but
also mouse strain-dependent.
When the OE and underlying tissue is removed from
the nasal cavity, it can be inserted into a small aperture
Ussing chamber for ion transport studies [77]. In all five CF
mutant strains tested, we observed a significantly increased
(P < 0.01) Isc response to amiloride ( Isc amil) (Fig. 7). This
includes the Cftrtm1Eur F508del mutation on different genetic
backgrounds and the Cftrtm1Cam knockout allele on mixed
background (C57Bl/6-129) and FVB. An enhanced Isc amil
and PDamil is also observed in human CF airway epithelia,
supporting the hypothesis that CF airway disease results, at
least in part, from ENaC hyperactivity. Consequently, ENaC
is a primary therapeutic target in CF mutant mice [15,78].
In excised nasal OE from Cftrtm1Eur F508del mice, Cftr
mRNA expression is normal (Scholte et al., data not shown),
whereas CFTR protein expression is much reduced (Fig. 6A
and B) and in Cftrtm1Cam knockout mice this signal is absent
(not shown). The absence of olfactory marker proteins from
CFTR-expressing cells argues that CFTR is not expressed
in ciliated sensory neurons, but in a subset of microvillous
secondary chemosensory cells (named brush cells), where it
is co-localized with ENaC (De Jonge et al, unpublished data).
Whether this cell type represents a cell specialized for fluid
and electrolyte transport or plays a role in CFTR-dependent
regeneration of the olfactory system [79,80] remains to be
established.
Forskolin responses observed in different mouse strains are
complex parameters that cannot, at face value, be attributed
to CFTR activity. In Cftrtm1Cam C57Bl/6-129 knockout mice,
we observe a forskolin response comparable to normal
littermates. However, in Cftrtm1Cam FVB mutant mice this
response is virtually absent (Fig. 7). These data suggest that
a cAMP-stimulated Cl− conductance other than CFTR is
present in the OE of Cftrtm1Cam C57Bl/6-129 mice, but is
absent from Cftrtm1Cam FVB mice. Studies are underway to
identify this cAMP-stimulated Cl− conductance.
In all Cftrtm1Eur F508del backcrossed strains the nasal
forskolin-stimulated Isc in mutant mice is comparable to
normal littermates. Because this includes the FVB strain
10. M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171 S161
Fig. 6. CFTR immunostaining in mouse olfactory epithelium (OE) and trachea. In wild-type FVB (FVB WT) mice punctuated CFTR staining in brush cells of
the olfactory epithelium (OE) (A) and diffuse CFTR staining in ciliated cells of the trachea (C) is observed. In homozygous mutant littermates (FVB F508del
Cftrtm1Eur), the CFTR staining in brush cells of the OE (B) and ciliated cells of the trachea (D) and is much reduced. The CFTR signal is completely absent in
sections from Cftrtm1Cam FVB knockout mice, stained in parallel (not shown).
that does not show a response with the Cftrtm1Cam knockout
allele, one possible explanation is that the nasal forskolin-
stimulated Isc reflects residual CFTR activity resulting from
partial processing of the mutant protein [9]. However, the
nasal forskolin-stimulated Isc does not influence the enhanced
amiloride response in this mutant strain (Fig. 7). Together
with the immunohistology data (Fig. 6A, B), this argues that
there is not a linear relationship between the nasal forskolin-
stimulated Isc and the number of active CFTR channels.
4.10. Submucosal glands in CF mutant mice
Mice have composite serous and mucous submucosal
glands, not only in the nasal cavity, but also in the upper
part of the trachea. This is relevant, because abnormal
submucosal glands are thought to play an important role
in the development of CF airway disease, contributing to
fluid hypo-secretion and the abnormal consistency of mucus
in the proximal airways [81,82]. Secretion of fluid from
submucosal glands is dependent on two complementary
combinations of transport systems, in particular basolateral
K+ channels and apical CFTR Cl− channels, regulated by
the intracellular calcium and cAMP signaling pathways,
respectively. The cAMP-stimulated component is lacking
in tracheal glands from CF mutant mice, and is therefore
thought to be CFTR-dependent [81,82]. The contribution
of CFTR-mediated secretion by submucosal glands to CF
mouse pathology is unclear, but this experimental paradigm
constitutes a valid model to study CFTR function and test
drugs targeting CFTR.
It is important to note that the submucosal glands of the
nasal cavity, and their contribution to its CF phenotype, have
not yet been studied. It is conceivable, that nasal submucosal
cells have properties different from tracheal glands, since they
have developed in a different cellular context.
4.11. CFTR expression in the mouse trachea
The cellular architecture of the pseudostratified columnar
mouse tracheal epithelium is similar to that of the nasal
airways. It is dominated by CFTR-expressing ciliated cells
(Fig. 6C and D), interspersed with infrequent mucus secreting
neuroendocrine and Clara cells. Mucociliary clearance and
epithelial ion transport are not significantly affected in tracheal
11. S162 M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171
Fig. 7. Summary of short-circuit current (Isc) measurements in nasal epithelium. Short-circuit current responses ( Isc) to amiloride (10 μM) and forskolin
(10 μM) were measured in isolated nasal epithelium in an Ussing chamber as described in [77]. Bioelectric characteristics of 12–18 weeks old normal (nn)
and homozygous mutant (dd) mice from the different backcrossed Cftrtm1Eur F508del strains (FVB, FVB/129, C57Bl/6, 129sv) are compared. Homozygous
Cftrtm1Cam knockout mice were analysed in FVB (FVB/n -/-), and the original Cftrtm1Cam strain (C57Bl/6-129 -/-), the latter compared to their normal (nn)
littermates. The number of individual animals measured per experimental group is given in parenthesis. The amiloride response is significantly (P < 0.01)
increased in all Cftr mutant strains compared to their wild-type littermates, except for the 129sv backcross. The forskolin response does not differ significantly
between the mutant strains and the wild-type mice in this assay, with the exception of the FVB Cftrtm1Cam knockout mouse strain (P < 0.001).
epithelia from CFTR knockout mice [83], but see [84].
Interestingly, in tracheal epithelia the Ca2+-activated Cl−
channel (CaCC) is the dominant apical membrane Cl−
channel, CFTR activity is only observed after maximal
stimulation of CaCC [85].
4.12. CaCC: the dark horse in CF pathology
CaCC is expressed in the apical membrane of epithelia
affected by CF, where it operates in parallel with CFTR, but
is regulated by a different intracellular signaling pathway.
As a result, CaCC is a potentially important element in CF
pathology. The recent demonstration that the TMEM16A gene
encodes a CaCC [86–88] will allow the molecular behaviour
of CaCC in different secretory epithelia to be elucidated.
Interestingly, TMEM16A knockout mice exhibit phenotypic
abnormalities consistent with a role in fluid and electrolyte
homeostasis [64–66]. It is therefore likely that the phenotype
of both CF mutant mice and CF patients is, at least in part,
dependent on TMEM16A expression and function. To what
extent CaCC affects the phenotypic abnormalities observed in
different organs of CF mice, as well as in CF patients, remains
to be established.
4.13. CFTR expression in the mouse lung
In mouse lung, Cftr mRNA expression is readily detected
by RT-PCR and in situ hybridization [89], but expression
levels are 10-fold lower than in nasal epithelium (Scholte et
al., not shown). Immunohistological detection of CFTR ex-
pression in the lung, confirmed by using CFTR knockout mice
as a negative control, is not feasible with current methods.
Mouse bronchi possess ciliated and mucus producing cells in
a declining proximal to distal distribution, whereas airways
in the mouse lung are dominated by Clara cells, constituting
60–80% of the total epithelial cell population. Clara cells are
non-ciliated secretory cells that (i) display phenotypic adapta-
tion and proliferative capacity during injury and inflammation,
(ii) are capable of adopting a mucus secretory phenotype [90–
92] and (iii) express functional CFTR [93,94]. Clara cells are
also prominent in human bronchioles, which, small in size,
but large in number, comprise a major portion of the airway
surface in the human lung. Like airways in the mouse lung,
human bronchioles lack cartilage and submucosal glands.
Of note, bronchioles play an important role in the initiation
of CF lung disease. For example, Tiddens et al. [95] and
Stick et al. [96] observed severe distal airway disease in at
least 50% of CF patients before the age of 5 years, including
patients free of bacterial colonization. Notwithstanding impor-
tant functional differences between human and murine airway
epithelia, murine nasal airway epithelia is therefore struc-
turally and functionally related to human bronchial airways,
whereas murine bronchi are similar to human bronchioles.
In alveolar epithelia, CFTR is expressed in type II epithelial
cells [97], where it is involved in cAMP-stimulated fluid
reabsorption [98]. Using an ingenious optical method, Lindert
et al. [99] demonstrated that basal fluid secretion in the
alveolar space was absent in Cftrtm1Unc homozygous mutant
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mouse lung and sensitive to the CFTR inhibitor CFTRinh-172
in normal mice.
5. Lung pathology in CF mutant mice
CF lung pathology has been studied intensively using CF
mutant mice and has been thoroughly reviewed [1–3]. Here
we present a summary of the literature and some relevant
recent data. Two issues are prominent in this field. First, to
what extent can we compare lung pathology between humans
and mice? Second, which parameters in a mouse model
provide relevant information about CF pathology?
In view of the major differences in size, cellular archi-
tecture, physiology, host-pathogen interactions, lifestyle and
lifespan, it is not surprising that CF mice do not reproduce
exactly the lung disease typical of CF patients. However, there
is considerable heterogeneity in the severity and progression
of CF lung disease in humans. Another caveat is that we know
surprisingly little about the early development of CF lung
disease from direct observations of human airways in situ,
especially the distal airways [95]. It is important to take these
considerations into account when comparing lung pathology
in mouse models and CF patients.
Several groups reported an inflammatory lung phenotype
associated with progressive tissue remodeling in different
unchallenged CF mutant mice [55,100–103]. The severity and
presentation of this phenotype, most prominent in a C57Bl/6
backcross of the Cftrtm1Unc mouse [19], appears to be strain-
and age-dependent. As discussed above, this phenotype will
be influenced by animal husbandry.
Here, we show that the inflammatory score based on quan-
titative histology is moderately, but significantly increased in
adult (16–19 weeks) unchallenged Cftrtm1Eur F508del (FVB)
mice kept in ventilated cages under pathogen-free conditions
(Fig. 8). Inflammation, enhanced levels of the macrophage at-
tractant CCL2/MCP-1 and increased numbers of macrophages
were reported in lung lavage fluid of a different colony of
Cftrtm1Eur F508del (129/FVB) mice, which were reduced
by the macrolide azythromycin [55]. Importantly, this in-
flammatory phenotype is not related to detectable pathogen
colonization, suggesting that CFTR deficiency by itself is
sufficient to generate a pro-inflammatory environment in the
mouse lung. At present it has not been established whether
this inflammatory lung phenotype is exclusively attributed to
the abnormal behaviour of cells from the myeloid lineage, in
particular macrophages [57,104], or whether abnormal proin-
flammatory signaling in CFTR-deficient epithelial cells plays
a role [104]. However, it should be noted that these animals
are not kept in a sterile environment. Exposure to fecal
bacteria, dust and other allergens are expected to be more
challenging to CF mutant animals than to normal littermates.
Therefore, it might be misleading to call the inflammatory
lung phenotype “spontaneous”. Moreover, animal husbandry,
genetic background and gender likely affect the data, which
explains why reports from different laboratories differ with
respect to the severity and presentation of the inflammatory
lung phenotype.
Fig. 8. Lung inflammatory score is increased in Cftrtm1Eur FVB mutant
mice. Paraffin sections were made from paraformaldehyde inflated lungs of
FVB Cftrtm1Eur F508del mice, homozygous normal (+/+, n = 5) and mutant
(dd, n = 6), age-matched (16–19 weeks comparable numbers of males and
females) littermates. The animals were kept in the Erasmus MC animal
facility, pathogen-free (FELASA+), in sterilized ventilated cages, provided
ad lib with normal chow and sterilized acidified water. Five micron sections
were stained with hematoxylin/eosin and scored blind by an independent
pathologist, for oedema, haemorrhages, recruitment of poly-morphonuclear
cells, macrophages, lymphocytes and plasma cells, lesions of alveolitis and
bronchitis, area of focal or diffuse consolidation, necrosis and metaplasia of
pneumonocytes. The scores in each category are expressed and averaged on
a scale of zero (no abnormalities) through one (mild inflammation) to five
(severe, destruction of the lung with lesions of alveolitis, severe bronchitis,
with necrosis and large areas of consolidation extended to all lobules). The
results show mild, but significant inflammation in CF mutant mice compared
to normal, under these conditions.
5.1. A challenged mouse counts for two
Waiting for CF mice to develop lung disease under stan-
dard laboratory conditions is a tedious, ineffective method.
Therefore, CF mutant mice of various strains have been
subjected to bacterial challenge protocols, resulting in a
substantial, albeit bewildering, body of data.
CF mice were infected with Pseudomonas aeruginosa
(PA), because this opportunistic pathogen causes life threaten-
ing chronic infections in CF patients. At first sight this seems
a rather naïve approach, since host pathogen interactions are
very complex, differ in mice and humans and are certainly
not dependent on CFTR activity alone. Moreover, PA adapts
rapidly to different environments, from hospital taps and
shower heads to patient lungs, forming mucoid variants and
biofilms. Nevertheless, several authors report enhanced mor-
tality and reduced clearance of acute PA infection in loss of
function CF mutants [105–108] and G551D mice [109]. What
causes mortality in acute PA infection models is not obvious,
and should probably be distinguished from the intensity and
resolution of inflammation. Interestingly, epithelial, but not
macrophage, specific Cftr gene complementation is necessary
and sufficient to protect G551D CF mice from PA mortality
[110].
While on the whole data from acute PA infection experi-
ments are satisfactory, several concerns need to be addressed.
It proved difficult to establish a true chronic infection with
clinical PA isolates. Mice, including CF mice, either rapidly
clear an intranasal or intra-tracheal dose of PA or they die
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quickly depending on the dose and method of delivery. Dif-
ferent clinical isolates and delivery methods have therefore
been tested [111,112]. Virulent PA strains embedded in agar
beads elicit a prolonged inflammatory reaction, simulating
chronic infection. Interestingly, responses to this challenge
are more severe in CF mutant mice [113], including the gut-
corrected version of the Cftrtm1Unc knockout mouse [114].
In the latter model, azithromycin reduced inflammation and
improved bacterial clearance after a challenge with alginate
embedded PA [115]. Though this is an elegant way to create
a prolonged inflammatory challenge, it does not necessarily
reflect accurately the response of human CF epithelia to PA
infection or how biofilms are formed and maintained in CF
lungs.
In a different approach, Pier and coauthors showed that
delivery of PA in drinking water results in low level chronic
colonization of CF mouse airways, but not those of controls
[116,117]. In this model, the inability to clear infection
was linked to abnormal IL-1β signaling in epithelial cells
[118]. However, this model lacked the intense inflammatory
responses of mice infected with high doses of PA.
Intra-tracheal lipopolysaccharide (LPS) isolated from bac-
teria can also be used to induce transient lung inflammation.
Similar to bacterial infection, this response is characterized
by activation of alveolar macrophages, cytokine production,
a massive neutrophil influx, but little or no mortality. The
responses seen in these models are primarily triggered by Toll
like receptors (TLR) of the innate immune system. In addition
to TLR2 and TLR3 responses to LPS, bacterial flagellin is
recognized by TLR5 and can be used to provoke an inflam-
matory response [119,120]. In CF mutant mice responses to
LPS challenge are stronger than in normal mice [56,57].
5.2. Mucus production in CF mutant mouse lung
Mucus plugging of the airways is an important aspect of CF
lung disease. This has been attributed to abnormal hydration
of CF mucus by the surface epithelium hyperabsorbing Na+
[78] and alternatively to a lack of bicarbonate secreting
cells, which prevents normal mucus hydration and secretion
[33,34,121]. Furthermore, goblet cell hyperplasia and mucus
hyper-secretion induced by inflammation are likely to play a
role.
In the CF mouse nasal cavity, goblet cell hyperplasia
does not correlate with infection [75]. In distal airways of
the normal mouse lung, we observe no goblet cells using
PAS/Alcian Blue, Muc5AC, and Clca3/Gob5 stains. However,
in proximal airways of the lung, second and third bifurcation
bronchi, we do observe areas containing PAS/Alcian Blue and
Clca3/Gob-5 positive cells.
Kent et al. [101] observed increased PAS staining together
with other lung alterations in unchallenged congenic C57Bl/6
Cftrm1Unc mice. In an independent study, Durie et al. [19]
confirmed progressive lung disease and airway accumula-
tion of mucus in the same mutant strain. Cftrtm1Hgu mice
demonstrate mucus accumulation compared to controls after
challenge with bacteria [107]. By contrast, Cressman et al.
[122] reported that Cftrtm1Unc mice with different genetic
backgrounds did not exhibit goblet cell hyperplasia com-
pared to normal controls, and both groups of mice responded
similarly to ovalbumin challenge. More recently, Touqui and
colleagues observed increased numbers of mucus producing
cells and MUC5AC antigen in lungs of homozygous mu-
tant C57Bl/6 Cftrtm1Unc mice [123]. Of note, the authors
demonstrated that this response was mediated by abnormal
activation of cytosolic phospholipase A2 and could be reduced
by a specific inhibitor of this enzyme [123]. Similar results
were obtained in Cftrtm1Eur mice (Scholte et al., unpublished
results).
In summary, unchallenged CF mutant mice present a mod-
est, but significant, increase in the number of mucus producing
cells in the proximal airways and mucus hypersecretion in
lavage. However, overt airway plugging and air trapping such
as that reported in CF patients and recently in CF ferrets and
pigs is not observed. Because cells that stain with standard
goblet cell markers are confined to the proximal airways in the
mouse lung, careful analysis of sections is required. In view
of the known susceptibility of this parameter to environmental
and genetic factors care must be taken to provide properly
controlled conditions.
The molecular biology of this response remains to be
investigated. On the basis of available data it is most likely
an adaptive response of Clara cells to the pro-inflammatory
environment created by CFTR deficiency. Whether a similar
response occurs in human distal airways is not known.
Finally, newborn mice overexpressing the β subunit of
ENaC exhibit a considerably stronger version of the mucus
hypersecretion phenotype (for further information see the
accompanying article [17]).
6. Bioactive lipids and CF lung disease
CFTR mutant mice display distinct abnormalities in the
metabolism of several lipids that are known to play a
role in intra- and extracellular signaling of inflammation
and apoptosis. Freedman et al. [124,125] first reported that
Cftrtm1Unc C57Bl/6 knockout mice show an increased ratio
of arachidonic acid (AA) to docosahexaenoic acid (DHA),
which was reversed by oral DHA. Importantly, this finding
correlated with a normalization of several phenotypic param-
eters in CF mutant mice [126]. More recently, Radzioch and
colleagues confirmed the abnormal AA/DHA ratio in adult
Cftrtm1Unc C57Bl/6 CF mutant mice [101] and CF patients
[127]. Furthermore, these authors reported reduced levels of
major ceramide species in plasma and tissue samples in both
species, which were reversed by treatment with the vita-
min A analogue fenretinide (N-(4-hydroxyphenyl)retinamide;
4-HPR) [127]. In CF mutant mice, fenretinide reduced the
sensitivity to intranasal challenge with PA [103] and reduced
bone abnormalities in CF knockout mice [128]. Whether
fenretinide can reduce the development of progressive CF
lung disease in patients remains to be established. Also the
molecular mechanism explaining the relationship between this
aspect of the CF phenotype and CFTR deficiency remains
14. M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171 S165
to be determined. It is hypothesized that both oral DHA
and fenretinide act by reducing pro-inflammatory signaling
through the PPAR pathway. In this context, Harmon et al.
[129] recently demonstrated that Cftrtm1Unc (B6.129P2) ho-
mozygous knockout mice (males and females, four weeks
old) have attenuated PPARγ activity due to reduced levels of
PPAR agonists, including 15-keto-PGE2. The authors further
showed that the PPARγ agonist rosiglitazone, used to treat
diabetes, corrected inflammatory and intestinal pathology in
the mouse model [129]. In accordance with these observa-
tions, we have reported previously that expression of the
15-keto-PGE2 precursor, PGF2α and Cbr2 (lung carbonyl
reductase), the enzyme responsible for PGF2α production in
airway epithelial cells, are reduced in BALF of homozygous
Cftrtm1Eur lungs [130]. How these findings relate to defective
CFTR function is not yet fully understood. Borot et al. [131]
presented evidence suggesting that CFTR can form a com-
plex with cytosolic phospholipase A2 through the S100A10
adaptor protein, which might explain the hyperactivity of
cellular phospholipase A2 in CF mutant mice and its effect on
mucin secretion [123]. We hypothesize that CFTR dysfunc-
tion affects the enzymatic pathways involved, either directly
or indirectly, through an adaptive process.
In a different CF mouse model carrying the Cftrtm1Hgu
partially active knockout allele (Table 1), Gulbins and col-
leagues reported an age-dependent accumulation of ceramide
species in lung tissue, which correlates with progressive lung
inflammation and sensitivity to PA challenge [102]. Reduction
of acid sphingomyelinase activity (ASM), either by mutation
(Smpd−/−) or using the ASM inhibitor amitryptilin normal-
ized ceramide levels and reduced lung pathology in these CF
mutant mice [102]. Further studies confirmed and extended
these data in the mouse model [132]; initial experiments in
CF patients demonstrated that amitriptylin is safe and might
be effective in reducing CF disease [133]. At this time,
the apparent contradictory changes in ceramide levels in CF
mutant mice between the Radzioch and Gulbins groups have
not been resolved using verified experimental protocols. It is
possible that differences in experimental protocols may cause
a bias towards distinct subsets of ceramide species.
7. Pumping the pipeline, testing experimental therapies in
mouse models
A key application of CF mutant mice is to test experimental
therapeutics. Elsewhere in this issue the development of
pharmacotherapy [134] and gene therapy for CF [135] are
reviewed. New antagonists of ENaC have been developed and
are being tested in a mouse model of ENaC hyperactivity
[17]. Compounds affecting bioactive lipid metabolism and
downstream transcription factor activity were discussed in
the previous section. The cloning of the CaCC TMEM16A
also presents new opportunities for intervention. As discussed
below, several experimental therapeutics have been tested in
CF mouse models.
7.1. CFTR activators
The mutant strains bearing F508del and G551D were made
specifically to test compounds that aim to correct defec-
tive processing and trafficking (CFTR correctors) or defective
channel gating (CFTR potentiators). A caveat is that these mu-
tations are introduced into the mouse CFTR sequence, which
differs from the human sequence. Therefore, the properties of
the mutant protein, including its interaction with therapeutic
drugs, might differ from the human form. Indeed, the murine
CFTR Cl− channel exhibits significant differences in its bio-
physical properties and regulation compared to human CFTR.
These differences include (i) a smaller single-channel conduc-
tance, (ii) a markedly different pattern of channel gating and
(iii) insensitivity to AMP-PNP and pyrophosphate (PPi), two
agents that potentiate robustly the activity of human CFTR
[136–138]. Consistent with these data, a number of CFTR
potentiators (e.g. VRT-532, NPPB-AM and VX-770), which
augment strongly human CFTR channel gating are without
effect on murine CFTR [139,140]. Moreover, Scott-Ward et
al. [141] demonstrated that transfer of both nucleotide-binding
domains (NBDs) of murine CFTR to human CFTR endowed
the human CFTR Cl− channel with the gating behaviour and
pharmacological properties of the murine CFTR Cl− chan-
nel. These data suggest that human-murine CFTR chimeras
might be employed to identify the binding sites of CFTR
potentiators.
7.2. CFTR correctors
Because CFTR correctors target the cellular protein pro-
cessing and trafficking machinery, they are less dependent
on a direct interaction with the CFTR protein. As a re-
sult, a number of CFTR correctors have been successfully
tested in the F508del mouse model (De Jonge et al., in
preparation). For example, miglustat, currently under eval-
uation as a therapy for Gauchers disease, shows limited,
but significant, efficacy in the Cftrtm1Eur mouse model by
reducing the amiloride response in the nasal epithelium
[142] and increasing F508del CFTR processing efficiency in
intestinal epithelium [143]. Moreover, intraperitoneal instil-
lation of sildenafil and vardenafil, related phosphodiesterase
5 inhibitors, activated nasal Cl− conductance in Cftrtm1Eur
F508del mice, but not Cftrtm1Cam null mice with the same
genetic background [144]. In a recent study, Robert et al.
[145] showed that glafenine, an off-patent drug with analgesic
properties, improved the trafficking of human F508del-CFTR
in BHK and CFBE41o− cells to 40% of normal values.
In isolated intestinal epithelia from Cftrtm1Eur mice, glafe-
nine increased the Isc response to forskolin plus genistein,
and it partially restored salivary secretion in vivo in mutant
mice, without causing adverse effects [145]. These results
illustrate that mutant CFTR mouse strains can play a signif-
icant role in testing experimental drugs that are identified in
high-throughput screening programs [134].
15. S166 M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171
7.3. Anti-inflammatory drugs
In addition to drugs targeting the bioactive lipid pathways
(see above, section 6), anti-inflammatory drugs can be investi-
gated successfully in mouse models. The macrolide antibiotic
azithromycin has anti-inflammatory action and reduced the
hyper inflammatory condition in Cftrtm1Eur F508del mutant
mice [55], apparently through an action at the level of alve-
olar macrophages [146], rather than at the level of epithelial
cells [104]. Similar results were reported for the Cftrtm1Unc-
TgN(FABPCFTR) gut corrected CFTR null mouse model
challenged with a clinical PA isolate [115]. The synthetic
triterpenoid CDDO reduced the hyper inflammatory pheno-
type of a R117H mutant mouse model, presumably through
inhibition of the NF-κB pro-inflammatory pathway [147].
8. Modifier genes and new therapeutic targets
Not only environmental factors but also genetic elements
determine CF pathology. First, the many different mutations
in the CFTR gene found in CF patients give rise to variable
levels of residual activity [27,148]. A different category of
genetic variations, often referred to as “modifier genes”,
are polymorphisms in genes that affect CFTR function or
pathways involved in CF pathology [149–151]. Identification
of modifier genes not only improves the CF diagnosis, but
also identifies novel therapeutic targets. CF mouse models
and the toolbox allowing genetic experiments in this species
offer unique opportunities to study modifier genes and their
potential therapeutic effects. First, phenotypic differences
between strains in different genetic backgrounds can be
analyzed [152,153] followed by gene mapping [154–156].
Current developments in gene and transcriptome sequencing
technology (“deep sequencing”) are likely to boost this field.
Another approach is to create specific mutations in candidate
genes that emerge from genetic or physiological studies in the
CF patient population. As an example, the identification of the
neutrophil marker IFRD1 as a modifier of lung disease in CF
patients was supported by analysis of Ifrd1 mutants in mice
[58]. Moreover, using KCNE3 knockout mice, Preston et al.
[157] recently demonstrated that the K+ channel β subunit
KCNE3 might act as a modifier gene in CF. Heteromultimeric
KCNQ1/KCNE3 K+ channels provide the driving force for
Cl− exit across the apical membrane of airway and intestinal
epithelia. In KCNE3 knockout mice, CFTR-mediated Cl−
secretion is attenuated markedly [157]. The availability of an
increasing number of conditional and inducible mutant mouse
strains, allows further validation of putative target genes. The
EUCOMM program (http://www.eucomm.org/), which aims
to generate conditional mutant embryo stem cell lines of
all functional genes in the mouse genome is linked to the
EUMODIC program (http://www.eumodic.org/aboutus.html)
which coordinates phenotypic analysis of derived strains.
Crossbreeding of selected mutations with mutant Cftr alleles
is expected to improve our knowledge of CF pathology.
9. Conclusion and perspective
We can conclude that despite their obvious limitations,
Cftr mutant mouse models have delivered. First, a better
understanding of CF pathology beyond clinical and cell
culture studies. Second, novel and promising therapeutics
have been identified, which are now being tested in these
models. Large animal models will certainly offer exciting new
dimensions in this field. However, the versatility and relative
affordability of mouse strains in genetic and physiological
studies will ensure their continued use.
Acknowledgements
We thank EuroCareCF (LSHM-CT-2005-018932), the
Dutch CF Foundation NCFS (R.B-O.) and the Dutch Maag
Lever Darm Stichting MLDS (M.W.), for financial support.
We gratefully acknowledge the expert assistance of Michel
Huerre (Institut Pasteur, Paris, France) in the quantitative
analysis of histological data.
Conflict of interest
All authors report no conflict of interest
References
[1] Grubb BR, Boucher RC. Pathophysiology of gene-targeted mouse
models for cystic fibrosis. Physiol Rev 1999;79(1 Suppl):S193–214.
[2] Scholte BJ, Davidson DJ, Wilke M, De Jonge HR. Animal models of
cystic fibrosis. J Cyst Fibros 2004;3(Suppl 2):183–90.
[3] Guilbault C, Saeed Z, Downey GP, Radzioch D. Cystic fibrosis mouse
models. Am J Respir Cell Mol Biol 2007;36:1–7.
[4] Rogers CS, Hao Y, Rokhlina T, et al. Production of CFTR-null
and CFTR- F508 heterozygous pigs by adeno-associated virus-
mediated gene targeting and somatic cell nuclear transfer. J Clin
Invest 2008;118:1571–7.
[5] Rogers CS, Abraham WM, Brogden KA, et al. The porcine lung as
a potential model for cystic fibrosis. Am J Physiol Lung Cell Mol
Physiol 2008;295:L240–63.
[6] Sun X, Yan Z, Yi Y, et al. Adeno-associated virus-targeted disruption
of the CFTR gene in cloned ferrets. J Clin Invest 2008;118:1578–83.
[7] Meyerholz DK, Stoltz DA, Pezzulo AA, Welsh MJ. Pathology of
gastrointestinal organs in a porcine model of cystic fibrosis. Am J
Pathol 2010;176:1377–89.
[8] Stoltz DA, Meyerholz DK, Pezzulo AA, et al. Cystic fibrosis pigs
develop lung disease and exhibit defective bacterial eradication at
birth. Sci Transl Med 2010;2:29ra31.
[9] French PJ, van Doorninck JH, Peters RH, et al. A F508 mutation
in mouse cystic fibrosis transmembrane conductance regulator results
in a temperature-sensitive processing defect in vivo. J Clin Invest
1996;98:1304–12.
[10] van Doorninck JH, French PJ, Verbeek E, et al. A mouse model for
the cystic fibrosis F508 mutation. EMBO J 1995;14:4403–11.
[11] Delaney SJ, Alton EW, Smith SN, et al. Cystic fibrosis mice carrying
the missense mutation G551D replicate human genotype-phenotype
correlations. EMBO J 1996;15:955–63.
[12] Hodges CA, Cotton CU, Palmert MR, Drumm ML. Generation of a
conditional null allele for Cftr in mice. Genesis 2008;46:546–52.
[13] Mall M, Grubb BR, Harkema JR, O’Neal WK, Boucher RC. Increased
airway epithelial Na+ absorption produces cystic fibrosis-like lung
disease in mice. Nat Med 2004;10:487–93.
[14] Mall MA, Harkema JR, Trojanek JB, et al. Development of chronic
16. M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171 S167
bronchitis and emphysema in β-epithelial Na+ channel-overexpressing
mice. Am J Respir Crit Care Med 2008;177:730–42.
[15] Mall MA. Role of the amiloride-sensitive epithelial Na+ channel in
the pathogenesis and as a therapeutic target for cystic fibrosis lung
disease. Exp Physiol 2009;94:171–4.
[16] Zhou Z, Treis D, Schubert SC, et al. Preventive but not late amiloride
therapy reduces morbidity and mortality of lung disease in βENaC-
overexpressing mice. Am J Respir Crit Care Med 2008;178:1245–56.
[17] Zhou Z, Duerr J, Johannesson B, et al. The βENaC-overexpressing
mouse as a model of cystic fibrosis lung disease. J Cyst Fibros
2011;10(Suppl 2):172–82.
[18] Reynaert I, Van Der Schueren B, Degeest G, et al. Morphological
changes in the vas deferens and expression of the cystic fibro-
sis transmembrane conductance regulator (CFTR) in control, F508
and knock-out CFTR mice during postnatal life. Mol Reprod Dev
2000;55:125–35.
[19] Durie PR, Kent G, Phillips MJ, Ackerley CA. Characteristic mul-
tiorgan pathology of cystic fibrosis in a long-living cystic fibro-
sis transmembrane regulator knockout murine model. Am J Pathol
2004;164:1481–93.
[20] Wright JT, Kiefer CL, Hall KI, Grubb BR. Abnormal enamel de-
velopment in a cystic fibrosis transgenic mouse model. J Dent Res
1996;75:966–73.
[21] Chang HY, Mitzner W. Sex differences in mouse models of asthma.
Can J Physiol Pharmacol 2007;85:1226–35.
[22] Kent G, Oliver M, Foskett JK, et al. Phenotypic abnormalities in
long-term surviving cystic fibrosis mice. Pediatr Res 1996;40:233–41.
[23] Cottart CH, Bonvin E, Rey C, et al. Impact of nutrition on phenotype
in CFTR-deficient mice. Pediatr Res 2007;62:528–32.
[24] Werner A, Bongers ME, Bijvelds MJ, de Jonge HR, Verkade HJ. No
indications for altered essential fatty acid metabolism in two murine
models for cystic fibrosis. J Lipid Res 2004;45:2277–86.
[25] Bijvelds MJ, Bronsveld I, Havinga R, Sinaasappel M, de Jonge
HR, Verkade HJ. Fat absorption in cystic fibrosis mice is impeded by
defective lipolysis and post-lipolytic events. Am J Physiol Gastrointest
Liver Physiol 2005;288:G646–53.
[26] Ohlsson L, Hjelte L, Huhn M, et al. Expression of intestinal and lung
alkaline sphingomyelinase and neutral ceramidase in cystic fibrosis
F508del transgenic mice. J Pediatr Gastroenterol Nutr 2008;47:547–
54.
[27] Rowntree RK, Harris A. The phenotypic consequences of CFTR
mutations. Ann Hum Genet 2003;67:471–85.
[28] Thiagarajah JR, Verkman AS. CFTR pharmacology and its role in
intestinal fluid secretion. Curr Opin Pharmacol 2003;3:594–9.
[29] Valverde MA, Vazquez E, Munoz FJ, et al. Murine CFTR channel
and its role in regulatory volume decrease of small intestine crypts.
Cell Physiol Biochem 2000;10:321–8.
[30] Venkatasubramanian J, Ao M, Rao MC. Ion transport in the small
intestine. Curr Opin Gastroenterol 2010;26:123–8.
[31] Kunzelmann K, Mall M. Electrolyte transport in the mammalian
colon: mechanisms and implications for disease. Physiol Rev
2002;82:245–89.
[32] Leverkoehne I, Holle H, Anton F, Gruber AD. Differential expression
of calcium-activated chloride channels (CLCA) gene family members
in the small intestine of cystic fibrosis mouse models. Histochem Cell
Biol 2006;126:239–50
[33] Quinton PM. Birth of Mucus. Am J Physiol Lung Cell Mol Physiol
2009;298:L13-4.
[34] Garcia MA, Yang N, Quinton PM. Normal mouse intestinal mucus
release requires cystic fibrosis transmembrane regulator-dependent
bicarbonate secretion. J Clin Invest 2009;119:2613–22.
[35] Malmberg EK, Noaksson KA, Phillipson M, et al. Increased levels
of mucins in the cystic fibrosis mouse small intestine and modulator
effects of the Muc1 mucin expression. Am J Physiol Gastrointest
Liver Physiol 2006;291:G203–10.
[36] Charizopoulou N, Wilke M, Dorsch M, et al. Spontaneous rescue
from cystic fibrosis in a mouse model. BMC Genet 2006;7:18.
[37] Wilke M, Bot A, Jorna H, Rensen S, De Jonge H. Correction of
the dF508-CFTR processing defect by low temperature incubation of
mouse intestine. Pediatr Pulmonol Suppl 2003;36:196
[38] Best JA, Quinton PM. Salivary secretion assay for drug efficacy for
cystic fibrosis in mice. Exp Physiol 2005;90:189–93.
[39] Noël S, Strale PO, Dannhoffer L, et al. Stimulation of salivary
secretion in vivo by CFTR potentiators in Cftr+/+ and Cftr−/− mice. J
Cyst Fibros 2008;7:128–33.
[40] Catalán MA, Nakamoto T, Gonzalez-Begne M, et al. Cftr and ENaC
ion channels mediate NaCl absorption in the mouse submandibular
gland. J Physiol 2010;588:713–24.
[41] Jouret F, Bernard A, Hermans C, et al. Cystic fibrosis is associated
with a defect in apical receptor-mediated endocytosis in mouse and
human kidney. J Am Soc Nephrol 2007;18:707–18.
[42] Jouret F, Devuyst O. CFTR and defective endocytosis: new insights in
the renal phenotype of cystic fibrosis. Pflugers Arch 2009;457:1227–
36.
[43] Feranchak AP. Hepatobiliary complications of cystic fibrosis. Curr
Gastroenterol Rep 2004;6:231–9.
[44] Bodewes F, Wouthuizen Bakker M, Havinga R, Bijvelds M, de
Jonge H, Verkade HJ. Treatment with ursodeoxycholate, but not with
cholic acid, increases bile flow independently of cystic fibrosis trans-
membrane conductance regulator in mice. Pediatr Pulmonol Suppl
2007;30:376.
[45] Peters RH, van Doorninck JH, French PJ, et al. Cystic fibrosis
transmembrane conductance regulator mediates the cyclic adenosine
monophosphate-induced fluid secretion but not the inhibition of re-
sorption in mouse gallbladder epithelium. Hepatology 1997;25:270–7.
[46] Peters RH, French PJ, van Doorninck JH, et al. CFTR expression and
mucin secretion in cultured mouse gallbladder epithelial cells. Am J
Physiol Gastrointest Liver Physiol 1996;271:G1074–83.
[47] Divangahi M, Balghi H, Danialou G, et al. Lack of CFTR in skeletal
muscle predisposes to muscle wasting and diaphragm muscle pump
failure in cystic fibrosis mice. PLoS Genet 2009;5:e1000586.
[48] Vandebrouck C, Melin P, Norez C, et al. Evidence that CFTR is
expressed in rat tracheal smooth muscle cells and contributes to
bronchodilation. Respir Res 2006;7:113.
[49] Robert R, Savineau JP, Norez C, Becq F, Guibert C. Expression and
function of cystic fibrosis transmembrane conductance regulator in rat
intrapulmonary arteries. Eur Respir J 2007;30:857–64.
[50] Michoud MC, Robert R, Hassan M, et al. Role of the cystic fibrosis
transmembrane conductance channel in human airway smooth muscle.
Am J Respir Cell Mol Biol 2009;40:217–22.
[51] Di A, Brown ME, Deriy LV, et al. CFTR regulates phagosome
acidification in macrophages and alters bactericidal activity. Nat Cell
Biol 2006;8:933–44.
[52] Deriy LV, Gomez EA, Zhang G, et al. Diseasecausing mutations
in the cystic fibrosis transmembrane conductance regulator deter-
mine the functional responses of alveolar macrophages. J Biol Chem
2009;284:35926–38.
[53] Haggie PM, Verkman AS. Defective organellar acidification as a
cause of cystic fibrosis lung disease: reexamination of a recurring
hypothesis. Am J Physiol Lung Cell Mol Physiol 2009;296:L859–67.
[54] Barriere H, Bagdany M, Bossard F, et al. Revisiting the role of
cystic fibrosis transmembrane conductance regulator and counterion
permeability in the pH regulation of endocytic organelles. Mol Biol
Cell 2009;20:3125–41.
[55] Legssyer R, Huaux F, Lebacq J, et al. Azithromycin reduces sponta-
neous and induced inflammation in F508 cystic fibrosis mice. Respir
Res 2006;7:134.
[56] Meyer M, Huaux F, Gavilanes X, et al. Azithromycin reduces exag-
gerated cytokine production by M1 alveolar macrophages in cystic
fibrosis. Am J Respir Cell Mol Biol 2009;41:590–602.
[57] Bruscia EM, Zhang PX, Ferreira E, et al. Macrophages directly
contribute to the exaggerated inflammatory response in cystic fibrosis
transmembrane conductance regulator−/− mice. Am J Respir Cell
Mol Biol 2009;40:295–304.
[58] Gu Y, Harley IT, Henderson LB, et al. Identification of IFRD1 as a
17. S168 M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171
modifier gene for cystic fibrosis lung disease. Nature 2009;458:1039–
42.
[59] Kulka M, Gilchrist M, Duszyk M, Befus AD. Expression and
functional characterization of CFTR in mast cells. J Leukoc Biol
2002;71:54–64.
[60] Norkina O, Kaur S, Ziemer D, De Lisle RC. Inflammation of the
cystic fibrosis mouse small intestine. Am J Physiol Gastrointest Liver
Physiol 2004;286:G1032–41.
[61] Xu Y, Tertilt C, Krause A, Quadri LE, Crystal RG, Worgall S.
Influence of the cystic fibrosis transmembrane conductance regulator
on expression of lipid metabolism-related genes in dendritic cells.
Respir Res 2009;10:26.
[62] Bonvin E, Le Rouzic P, Bernaudin JF, et al. Congenital tracheal
malformation in cystic fibrosis transmembrane conductance regulator-
deficient mice. J Physiol 2008;586:3231–43.
[63] Liang H, Yang L, Ma T, Zhao Y. Functional expression of cystic
fibrosis transmembrane conductance regulator in mouse chondrocytes.
Clin Exp Pharmacol Physiol 2010;37:506–8.
[64] Rock JR, O’Neal WK, Gabriel SE, et al. Transmembrane protein
16A (TMEM16A) is a Ca2+-regulated Cl secretory channel in mouse
airways. J Biol Chem 2009;284:14875–80.
[65] Ousingsawat J, Martins JR, Schreiber R, Rock JR, Harfe BD, Kun-
zelmann K. Loss of TMEM16A causes a defect in epithelial Ca2+
-
dependent chloride transport. J Biol Chem 2009;284:28698–703.
[66] Huang F, Rock JR, Harfe BD, et al. Studies on expression and
function of the TMEM16A calcium-activated chloride channel. Proc
Natl Acad Sci U S A 2009;106:21413–8.
[67] Dif F, Marty C, Baudoin C, de Vernejoul MC, Levi G. Severe
osteopenia in CFTR-null mice. Bone 2004;35:595–603.
[68] Pashuck TD, Franz SE, Altman MK, et al. Murine model for cystic
fibrosis bone disease demonstrates osteopenia and sex-related differ-
ences in bone formation. Pediatr Res 2009;65:311–6.
[69] Haston CK, Li W, Li A, Lafleur M, Henderson JE. Persistent osteope-
nia in adult cystic fibrosis transmembrane conductance regulator-
deficient mice. Am J Respir Crit Care Med 2008;177:309–15.
[70] Bronckers A, Kalogeraki L, Jorna HJ, et al. The cystic fibrosis trans-
membrane conductance regulator (CFTR) is expressed in maturation
stage ameloblasts, odontoblasts and bone cells. Bone 2010;46:1188–
96.
[71] Weinert S, Jabs S, Supanchart C, et al. Lysosomal pathology and
osteopetrosis upon loss of H+ driven lysosomal Cl accumulation.
Science 2010;328:1401–3.
[72] Kornak U, Kasper D, Bosl MR, et al. Loss of the ClC-7 chloride
channel leads to osteopetrosis in mice and man. Cell 2001;104:205–
15.
[73] Paccou J, Zeboulon N, Combescure C, Gossec L, Cortet B. The
prevalence of osteoporosis, osteopenia, and fractures among adults
with cystic fibrosis: a systematic literature review with meta-analysis.
Calcif Tissue Int 2010;86:1–7.
[74] Sermet-Gaudelus I, Castanet M, Retsch-Bogart G, Aris RM. Update
on cystic fibrosis-related bone disease: a special focus on children.
Paediatr Respir Rev 2009;10:134–42.
[75] Hilliard TN, Zhu J, Farley R, et al. Nasal abnormalities in cystic
fibrosis mice independent of infection and inflammation. Am J Respir
Cell Mol Biol 2008;39:19–25.
[76] Grubb BR, Rogers TD, Kulaga HM, et al. Olfactory epithelia exhibit
progressive functional and morphological defects in CF mice. Am J
Physiol Cell Physiol 2007;293:C574–83.
[77] De Jonge HR, Ballmann M, Veeze H, et al. Ex vivo CF diagnosis by
intestinal current measurements (ICM) in small aperture, circulating
Ussing chambers. J Cyst Fibros 2004;3(Suppl 2):159–63.
[78] Boucher RC. Cystic fibrosis: a disease of vulnerability to airway
surface dehydration. Trends Mol Med 2007;13:231–40.
[79] Elsaesser R, Paysan J. Morituri te salutant? Olfactory signal transduc-
tion and the role of phosphoinositides. J Neurocytol 2005;34:97–116.
[80] Elsaesser R, Montani G, Tirindelli R, Paysan J. Phosphatidyl-inositide
signalling proteins in a novel class of sensory cells in the mammalian
olfactory epithelium. Eur J Neurosci 2005;21:2692–700.
[81] Ianowski JP, Choi JY, Wine JJ, Hanrahan JW. Substance P stimu-
lates CFTR-dependent fluid secretion by mouse tracheal submucosal
glands. Pflugers Arch 2008;457:529–37.
[82] Ianowski JP, Choi JY, Wine JJ, Hanrahan JW. Mucus secretion by
single tracheal submucosal glands from normal and cystic fibro-
sis transmembrane conductance regulator knockout mice. J Physiol
2007;580:301–14.
[83] Grubb BR, Jones JH, Boucher RC. Mucociliary transport determined
by in vivo microdialysis in the airways of normal and CF mice. Am J
Physiol Lung Cell Mol Physiol 2004;286:L588–95.
[84] Zahm JM, Gaillard D, Dupuit F, et al. Early alterations in airway
mucociliary clearance and inflammation of the lamina propria in CF
mice. Am J Physiol Cell Physiol 1997;272:C853–9.
[85] MacVinish LJ, Gill DR, Hyde SC, et al. Chloride secretion in the
trachea of null cystic fibrosis mice: the effects of transfection with
pTrial10-CFTR2. J Physiol 1997;499:677–87.
[86] Schroeder BC, Cheng T, Jan YN, Jan LY. Expression cloning of
TMEM16A as a calcium-activated chloride channel subunit. Cell
2008;134:1019–29.
[87] Caputo A, Caci E, Ferrera L, et al. TMEM16A, a membrane protein
associated with calcium-dependent chloride channel activity. Science
2008;322:590–4.
[88] Yang YD, Cho H, Koo JY, et al. TMEM16A confers receptor-activated
calcium-dependent chloride conductance. Nature 2008;455:1210–5.
[89] Rochelle LG, Li DC, Ye H, Lee E, Talbot CR, Boucher RC. Distribu-
tion of ion transport mRNAs throughout murine nose and lung. Am J
Physiol Lung Cell Mol Physiol 2000;279:L14–24.
[90] Evans CM, Williams OW, Tuvim MJ, et al. Mucin is produced by
clara cells in the proximal airways of antigen-challenged mice. Am J
Respir Cell Mol Biol 2004;31:382–94.
[91] Park KS, Korfhagen TR, Bruno MD, et al. SPDEF regulates goblet
cell hyperplasia in the airway epithelium. J Clin Invest 2007;117:978–
88.
[92] Davis CW, Dickey BF. Regulated airway goblet cell mucin secretion.
Annu Rev Physiol 2008;70:487–512.
[93] Kulaksiz H, Schmid A, Honscheid M, Ramaswamy A, Cetin Y. Clara
cell impact in air-side activation of CFTR in small pulmonary airways.
Proc Natl Acad Sci U S A 2002;99:6796–801.
[94] Chinet TC, Gabriel SE, Penland CM, et al. CFTR-like chloride
channels in non-ciliated bronchiolar epithelial (Clara) cells. Biochem
Biophys Res Commun 1997;230:470–5.
[95] Tiddens HA, Donaldson SH, Rosenfeld M, Pare PD. Cystic fibro-
sis lung disease starts in the small airways: can we treat it more
effectively? Pediatr Pulmonol 2010;45:107–17.
[96] Stick SM, Brennan S, Murray C, et al. Bronchiectasis in infants
and preschool children diagnosed with cystic fibrosis after newborn
screening. J Pediatr 2009;155:623–8 e1.
[97] Fang X, Song Y, Hirsch J, et al. Contribution of CFTR to apical-
basolateral fluid transport in cultured human alveolar epithelial type II
cells. Am J Physiol Lung Cell Mol Physiol 2006;290:L242–9.
[98] Matthay MA, Robriquet L, Fang X. Alveolar epithelium: role in lung
fluid balance and acute lung injury. Proc Am Thorac Soc 2005;2:206–
13.
[99] Lindert J, Perlman CE, Parthasarathi K, Bhattacharya J. Chloride-
dependent secretion of alveolar wall liquid determined by optical-
sectioning microscopy. Am J Respir Cell Mol Biol 2007;36:688–96.
[100] Thomas GR, Costelloe EA, Lunn DP, et al. G551D cystic fibro-
sis mice exhibit abnormal regulation of inflammation in lungs and
macrophages. J Immunol 2000;164:3870–7.
[101] Kent G, Iles R, Bear CE, et al. Lung disease in mice with cystic
fibrosis. J Clin Invest 1997;100:3060–9.
[102] Teichgraber V, Ulrich M, Endlich N, et al. Ceramide accumulation
mediates inflammation, cell death and infection susceptibility in cystic
fibrosis. Nat Med 2008;14:382–91.
[103] Guilbault C, De Sanctis JB, Wojewodka G, et al. Fenretinide corrects
newly found ceramide deficiency in cystic fibrosis. Am J Respir Cell
Mol Biol 2008;38:47–56.
[104] Gavilanes X, Huaux F, Meyer M, et al. Azithromycin fails to re-
18. M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171 S169
duce increased expression of neutrophil-related cytokines in primary-
cultured epithelial cells from cystic fibrosis mice. J Cyst Fibros
2009;8:203–10.
[105] van Heeckeren AM, Schluchter MD, Xue W, Davis PB. Response to
acute lung infection with mucoid Pseudomonas aeruginosa in cystic
fibrosis mice. Am J Respir Crit Care Med 2006;173:288–96.
[106] Heeckeren A, Walenga R, Konstan MW, Bonfield T, Davis PB,
Ferkol T. Excessive inflammatory response of cystic fibrosis mice
to bronchopulmonary infection with Pseudomonas aeruginosa. J Clin
Invest 1997;100:2810–5.
[107] Davidson DJ, Dorin JR, McLachlan G, et al. Lung disease in the
cystic fibrosis mouse exposed to bacterial pathogens. Nat Genet
1995;9:351–7.
[108] Stotland PK, Radzioch D, Stevenson MM. Mouse models of chronic
lung infection with Pseudomonas aeruginosa: models for the study of
cystic fibrosis. Pediatr Pulmonol 2000;30:413–24.
[109] McMorran BJ, Palmer JS, Lunn DP, et al. G551D CF mice display an
abnormal host response and have impaired clearance of Pseudomonas
lung disease. Am J Physiol Lung Cell Mol Physiol 2001;281:L740–7.
[110] Oceandy D, McMorran BJ, Smith SN, et al. Gene complementa-
tion of airway epithelium in the cystic fibrosis mouse is necessary
and sufficient to correct the pathogen clearance and inflammatory
abnormalities. Hum Mol Genet 2002;11:1059–67.
[111] Bragonzi A, Paroni M, Nonis A, et al. Pseudomonas aeruginosa
microevolution during cystic fibrosis lung infection establishes clones
with adapted virulence. Am J Respir Crit Care Med 2009;180:138–45.
[112] Moser C, Van Gennip M, Bjarnsholt T, et al. Novel experimental
Pseudomonas aeruginosa lung infection model mimicking long-term
host-pathogen interactions in cystic fibrosis. APMIS 2009;117:95–
107.
[113] van Heeckeren AM, Schluchter MD, Drumm ML, Davis PB. Role
of Cftr genotype in the response to chronic Pseudomonas aerugi-
nosa lung infection in mice. Am J Physiol Lung Cell Mol Physiol
2004;287:L944–52.
[114] van Heeckeren AM, Schluchter M, Xue L, et al. Nutritional effects on
host response to lung infections with mucoid Pseudomonas aeruginosa
in mice. Infect Immun 2004;72:1479–86.
[115] Tsai WC, Hershenson MB, Zhou Y, Sajjan U. Azithromycin increases
survival and reduces lung inflammation in cystic fibrosis mice. In-
flamm Res 2009;58:491–501.
[116] Schroeder TH, Reiniger N, Meluleni G, Grout M, Coleman FT, Pier
GB. Transgenic cystic fibrosis mice exhibit reduced early clearance
of Pseudomonas aeruginosa from the respiratory tract. J Immunol
2001;166:7410–8.
[117] Coleman FT, Mueschenborn S, Meluleni G, et al. Hypersusceptibility
of cystic fibrosis mice to chronic Pseudomonas aeruginosa oropha-
ryngeal colonization and lung infection. Proc Natl Acad Sci U S A
2003;100:1949–54.
[118] Reiniger N, Lee MM, Coleman FT, Ray C, Golan DE, Pier GB.
Resistance to Pseudomonas aeruginosa chronic lung infection re-
quires cystic fibrosis transmembrane conductance regulator-modulated
interleukin-1 (IL-1) release and signaling through the IL-1 receptor.
Infect Immun 2007;75:1598–608.
[119] Raoust E, Balloy V, Garcia-Verdugo I, Touqui L, Ramphal R, Chig-
nard M. Pseudomonas aeruginosa LPS or flagellin are sufficient to
activate TLR-dependent signaling in murine alveolar macrophages
and airway epithelial cells. PloS One 2009;4:e7259.
[120] Morris AE, Liggitt HD, Hawn TR, Skerrett SJ. Role of Toll-like
receptor 5 in the innate immune response to acute P. aeruginosa
pneumonia. Am J Physiol Lung Cell Mol Physiol 2009;297:L1112–9.
[121] Quinton PM. Cystic fibrosis: impaired bicarbonate secretion and mu-
coviscidosis. Lancet 2008;372:415–7.
[122] Cressman VL, Hicks EM, Funkhouser WK, Backlund DC, Koller
BH. The relationship of chronic mucin secretion to airway disease
in normal and CFTR-deficient mice. Am J Respir Cell Mol Biol
1998;19:853–66.
[123] Dif F, Wu YZ, Burgel PR, et al. Critical role of cytosolic phospholi-
pase a2α in bronchial mucus hyper-secretion in CFTR-deficient mice.
Eur Respir J 2010 Apr 22. [Epub ahead of print]
[124] Freedman SD, Katz MH, Parker EM, Laposata M, Urman MY, Al-
varez JG. A membrane lipid imbalance plays a role in the phenotypic
expression of cystic fibrosis in cftr−/− mice. Proc Natl Acad Sci U S
A 1999;96:13995–4000.
[125] Freedman SD, Shea JC, Blanco PG, Alvarez JG. Fatty acids in cystic
fibrosis. Curr Opin Pulm Med 2000;6:530–2.
[126] Beharry S, Ackerley C, Corey M, et al. Long-term docosahexaenoic
acid therapy in a congenic murine model of cystic fibrosis. Am J
Physiol Gastrointest Liver Physiol 2007;292:G839–48.
[127] Guilbault C, Wojewodka G, Saeed Z, et al. Cystic fibrosis fatty
acid imbalance is linked to ceramide deficiency and corrected by
fenretinide. Am J Respir Cell Mol Biol 2009;41:100–6.
[128] Saeed Z, Guilbault C, De Sanctis JB, et al. Fenretinide prevents
the development of osteoporosis in Cftr-KO mice. J Cyst Fibros
2008;7:222–30.
[129] Harmon GS, Dumlao DS, Ng DT, et al. Pharmacological correction
of a defect in PPAR-γ signaling ameliorates disease severity in Cftr-
deficient mice. Nat Med 2010;16:313–8.
[130] Carvalho-Oliveira IM, Charro N, Aarbiou J, et al. Proteomic analysis
of naphthalene-induced airway epithelial injury and repair in a cystic
fibrosis mouse model. J Proteome Res 2009;8:3606–16.
[131] Borot F, Vieu DL, Faure G, et al. Eicosanoid release is increased by
membrane destabilization and CFTR inhibition in Calu-3 cells. PloS
One 2009;4:e7116.
[132] Becker KA, Riethmuller J, Luth A, Döring G, Kleuser B, Gulbins
E. Acid sphingomyelinase inhibitors normalize pulmonary ceramide
and inflammation in cystic fibrosis. Am J Respir Cell Mol Biol
2010;42:716–24
[133] Riethmüller J, Anthonysamy J, Serra E, Schwab M, Döring G, Gul-
bins E. Therapeutic efficacy and safety of amitriptyline in patients
with cystic fibrosis. Cell Physiol Biochem 2009;24:65–72.
[134] Becq F, Mall MA, Sheppard DN, Conese M, Zegarra-Moran O.
Pharmacological therapy for cystic fibrosis: From bench to bedside. J
Cyst Fibros 2011;10(Suppl 2):S129–45.
[135] Conese M, Ascenzioni F, Boyd AC, et al. Gene and cell therapy for
cystic fibrosis: From bench to bedside. J Cyst Fibros 2011;10(Suppl
2):129–45.
[136] Sheppard DN, Welsh MJ. Structure and function of the CFTR chloride
channel. Physiol Rev 1999;79(1 Suppl):S23–45.
[137] Lansdell KA, Delaney SJ, Lunn DP, Thomson SA, Sheppard DN,
Wainwright BJ. Comparison of the gating behaviour of human and
murine cystic fibrosis transmembrane conductance regulator Cl chan-
nels expressed in mammalian cells. J Physiol 1998;508:379–92.
[138] Lansdell KA, Kidd JF, Delaney SJ, Wainwright BJ, Sheppard DN.
Regulation of murine cystic fibrosis transmembrane conductance reg-
ulator Cl channels expressed in Chinese hamster ovary cells. J Physiol
1998;512:751–64.
[139] Van Goor F, Hadida S, Grootenhuis PD, et al. Rescue of CF airway
epithelial cell function in vitro by a CFTR potentiator, VX-770. Proc
Natl Acad Sci U S A 2009;106:18825–30.
[140] De Jonge H, Wilke M, Bot A, Jorna H, Wang W, Kirk K, et
al. Differential response of mouse versus human CFTR to CFTR
potentiators. Pediatr Pulmonol Suppl 2007;30:208.
[141] Scott-Ward TS, Cai Z, Dawson ES, et al. Chimeric constructs endow
the human CFTR Cl channel with the gating behavior of murine
CFTR. Proc Natl Acad Sci U S A 2007;104:16365–70.
[142] Noël S, Wilke M, Bot AG, De Jonge HR, Becq F. Parallel im-
provement of sodium and chloride transport defects by miglustat
(n-butyldeoxynojyrimicin) in cystic fibrosis epithelial cells. J Pharma-
col Exp Ther 2008;325:1016–23.
[143] Norez C, Noel S, Wilke M, et al. Rescue of functional delF508-
CFTR channels in cystic fibrosis epithelial cells by the α-glucosidase
inhibitor miglustat. FEBS Lett 2006;580:2081–6.
[144] Lubamba B, Lecourt H, Lebacq J, et al. Preclinical evidence that
sildenafil and vardenafil activate chloride transport in cystic fibrosis.
Am J Respir Crit Care Med 2008;177:506–15.
![Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171
www.elsevier.com/locate/jcf
Mouse models of cystic fibrosis: Phenotypic analysis and
research applications
Martina Wilkea,1
, Ruvalic M. Buijs-Offermanb
, Jamil Aarbioub,2
, William H. Colledgec
,
David N. Sheppardd
, Lhousseine Touquie
, Alice Bota
, Huub Jornaa
, Hugo R. de Jongea
,
Bob J. Scholteb,
*
a Erasmus MC, Biochemistry Department, 3000 CA Rotterdam, The Netherlands
b Erasmus MC, Cell Biology Department, 3000 CA Rotterdam, The Netherlands
c Physiological Laboratory, Department of Physiology, Development and Neuroscience, University of Cambridge, Cambridge CB2 3EG, UK
d University of Bristol, School of Physiology & Pharmacology, Medical Sciences Building, University Walk, Bristol BS8 1TD, UK
e Unité de Défense Innée et Inflammation, Unité Inserm U.874, Institut Pasteur, 75015 Paris, France
Abstract
Genetically modified mice have been studied for more than fifteen years as models of cystic fibrosis (CF). The large amount of
experimental data generated illuminates the complex multi-organ pathology of CF and raises new questions relevant to human disease. CF
mice have also been used to test experimental therapies prior to clinical trials. This review recapitulates the major phenotypic traits of CF mice
and highlights important new findings including aberrant alveolar macrophages, bone and cartilage abnormalities and abnormal bioactive lipid
metabolism. Novel data are presented on the intestinal and nasal physiology of F508del-CFTR CF mice backcrossed onto different genetic
backgrounds. Caveats, and sources of variability including age, gender and animal husbandry, are discussed. Interspecies differences limit
comparison of lung pathology in CF mice to the human disease. The recent development of genetically modified pigs and ferrets heralds the
application of more advanced animal models to CF research and drug development.
© 2011 European Cystic Fibrosis Society. Published by Elsevier B.V. All rights reserved.
Keywords: Cystic fibrosis; Animal models; CFTR mice; Lung disease; Intestinal disease; F508del-CFTR
1. Introduction
The life expectancy of cystic fibrosis (CF) patients in-
creased from around 20 to 40 years over the past two decades,
mainly due to improved nutrition and treatment of chronic
lung disease. However, there is still an urgent unmet need to
improve the lifespan and quality of life of CF patients. The
complex multi-organ pathophysiology of secretory epithelia
in CF is not yet completely understood, and a treatment
that sufficiently corrects the primary defect remains to be
developed.
* Corresponding author: B.J. Scholte, PhD, Erasmus MC, Cell Biology
Department, PO Box 2040, 3000 CA Rotterdam, The Netherlands. Tel.: +31
10 7043205; fax: +31 10 7044743.
E-mail address: b.scholte@erasmusmc.nl (B.J. Scholte)
1 Present address: Erasmus MC, Clinical Genetics Department.
2 Present address: Galapagos NV, Leiden.
1569-1993/$ - see front matter © 2011 European Cystic Fibrosis Society. Published by Elsevier B.V. All rights reserved.
Because access to relevant patient tissues is limited, genet-
ically modified mice are widely used to study CF pathology
and experimental therapeutics. These models have proven to
be very useful, but they have important limitations, due to
obvious physiological differences between humans and mice
[1–3]. Recently, pig [4,5] and ferret [6] cystic fibrosis trans-
membrane conductance regulator (CFTR) knockout animals
were reported, opening an exciting new field. Both models
show dominant intestinal disease [6,7], with severe perinatal
lethality. There is also evidence for reduced clearance of
bacterial lung infection in a limited number of long-lived CF
pigs [8]. In both models, F508del-CFTR mutants are being
developed, which will facilitate the testing of novel therapeu-
tic strategies for CF. The challenge for the time to come will
be to manage intestinal disease in CF pigs and ferrets to allow
better understanding of early lung disease, which is still an
underexplored area of CF pathology. While we can expect](https://image.slidesharecdn.com/1-s2-130830013718-phpapp02/85/1-s2-0-s1569199311600209-main-1-320.jpg)
![M. Wilke et al. / Journal of Cystic Fibrosis Volume 10 Suppl 2 (2011) S152–S171 S153
great progress from these models, their application is limited
by the considerable practical and financial issues involved.
This paper will focus on CF mouse models, presenting an
update on new developments and novel data, in particular on
the properties of the mutant strains made available to the CF
community through the EuroCareCF project.
2. CF mutant mice, portrait of an unruly family
Shortly after the discovery of the CFTR gene, several
groups set out to generate mouse strains with mutations in the
Cftr locus using the homologous recombination technology
and mouse embryonic stem cells pioneered by Mario Capec-
chi, Martin Evans and Oliver Smithies who jointly received
the Nobel Prize for these innovations in 2007. Table 1 lists
the genetically modified mouse strains produced by different
groups. Details of mouse strain nomenclature and availability
can be found at the Mouse Genome Informatics website
(http://www.informatics.jax.org/). Table 2 summarizes briefly
the major phenotypic traits of CF mice in comparison with the
human disease.
Several loss of function, or “knock-out” models were made
by interrupting the mouse Cftr coding region with a selectable
marker. The Cftrtm1Hgu shows low level of residual activity
of wild-type CFTR and lower mortality, compared to the
other two frequently used knockout strains Cftrtm1Cam and
Cftrtm1Unc (Table 1).
In addition, common CF causing mutations were intro-
duced into the mouse CFTR amino acid sequence. Three
strains with the F508del mutation in the mouse Cftr gene
were generated, two of which carry an intronic insertion that
may interfere with gene transcription (Table 1). By contrast,
the Cftrtm1Eur allele was made by the two step “hit and
run” recombination procedure, which removes the selectable
marker. Therefore, this allele only contains the F508del mu-
tation and has normal transcription rates in all tissues [9,10].
All three strains have been successfully used in phenotypic
studies. The F508del mutation in the mouse Cftr context
causes severely reduced amounts of fully glycosylated CFTR
protein (band C) similar to the human form, and a phenotype
comparable to CFTR loss of function mutants [9]. In cultured
gallbladder epithelial cells from F508del mutant mice, the
number of active CFTR Cl− channels observed in patch-
clamp experiments is increased by incubating cells at low
temperature [9], suggesting a similar temperature-dependent
processing defect to the F508del mutation in human CFTR.
Moreover, the G551D mutation which disrupts channel gating
has the typical CF phenotype caused by loss of function
mutants [11]. The less severe phenotype of the F508del-
and G551D-CFTR models when compared to CFTR knock-
outs, particularly lower perinatal mortality due to intestinal
obstruction, has been attributed to residual CFTR activity.
To overcome the limitations imposed by the severe in-
testinal disease presented by many knockout strains, “gut
corrected” hybrid strains were created using a transgene ex-
pressing CFTR from a promoter specific for the intestinal
epithelium (Table 1).
The most recent and important addition to the CF mouse
family is a conditional (“loxed”) Cftr deletion that can be
created by cell specific expression of the cre recombinase [12].
This genetically modified mouse will allow the contribution
of different cell lineages to CF pathology to be elucidated.
Although not a CFTR mouse model, genetically modi-
fied mice overexpressing the β subunit of the epithelial Na+
channel (ENaC) are important because they possess a lung
phenotype resembling that of CF [13–16]. For further infor-
mation about this mouse model and its applications, see Zhou
et al. [17].
3. Mouse models, sources of variation and practical
considerations
CF mutant mice are generally much less fertile than normal
mice, which makes it difficult to breed them from homozy-
gous mutant pairs. This is partly due to frequent obstruction
of the male vas deferens [18,19]. However, homozygous fe-
males are also poor breeders. Occasionally mice can be bred
successfully from homozygous pairs. However, there is risk
of “genetic drift”, the selection of compensating mutations in
either the background or the Cftr allele, which affect pheno-
type in an unpredictable way. Breeding CF mutant mice from
heterozygous animals is therefore recommended strongly. Im-
portantly, this breeding strategy yields age- and sex-matched
normal littermates, which are mandatory control for most
experiments. Breeding requires considerable investment in
DNA genotyping of all neonates by ear or tail clips, the
identification of individual animals by chip, toe- or ear-clips
and a database to track individual animals. Genotyping is
primarily required to identify heterozygous breeders, because
homozygous mutants are easily recognized by their typical
white incisor teeth [20].
There is consensus about the most prominent phenotypic
abnormalities of different Cftr mutant mice. Yet, close com-
parison of published data from different laboratories shows
considerable variation and even controversy about the nature
and extent of lung disease. One obvious source of variation
is the different Cftr mutations, some of which are associated
with residual CFTR function (Table 1). Furthermore, Cftr
alleles have been bred onto different genetic backgrounds,
which may cause subtle differences in phenotype, presumably
because of the expression of different “modifier genes”. Com-
plete genomes of several laboratory mouse strains are now
available (for further information, see Sanger mouse genome
project, http://www.sanger.ac.uk/resources/mouse/genomes/).
This resource will provide a new and powerful tool to ana-
lyze phenotypic differences between CFTR mutant strains. It
will also facilitate comparative studies of modifier genes and
potential therapeutic targets in the human population.
Animal husbandry is a critical issue, especially when in-
vestigating innate adaptive responses (e.g. lung inflammation),
which are by their very nature designed to reflect external
conditions. This can cause differences between strains, con-
tribute to inter-individual variation, and lead to differences
in results between laboratories. Unfortunately, there is no](data:image/gif;base64,R0lGODlhAQABAIAAAAAAAP///yH5BAEAAAAALAAAAAABAAEAAAIBRAA7)