An immunocytochemical assay to detect human CFTR expression following gene transfer.

Abstract

To assess gene therapy treatment for cystic fibrosis (CF) in clinical trials it is essential to develop robust assays that can accurately detect transgene expression in human airway epithelial cells. Our aim was to develop a reproducible immunocytochemical assay for human CFTR protein which can measure both endogenous CFTR levels and augmented CFTR expression after gene delivery.
We characterised an antibody (G449) which satisfied the criteria for use in clinical trials. We optimised our immunocytochemistry method and identified G449 dilutions at which endogenous CFTR levels were negligible in CF samples, thus enhancing detection of transgenic CFTR protein. After developing a transfection technique for brushed human nasal epithelial cells, we transfected non-CF and CF cells with a clinically relevant CpG-free plasmid encoding human CFTR.
The optimised immunocytochemistry method gave improved discrimination between CF and non-CF samples. Transfection of a CFTR expression vector into primary nasal epithelial cells resulted in detectable RNA and protein expression. CFTR protein was present in 0.05-10% of non-CF cells and 0.02-0.8% of CF cells.
We have developed a sensitive, clinically relevant immunocytochemical assay for CFTR protein and have used it to detect transgene-expressed CFTR in transfected human primary airway epithelial cells.

Full text

Page 1

An immunocytochemical assay to detect human CFTR expression following
gene transfer
Heather Davidsona,h,*, Abigail Wilsona,h, Robert D. Graya,h, Alex Horsleya,h, Ian A. Pringlec,h,
Gerry McLachlanb,h, Angus C. Nairnf,g, Cordelia Stearnsf,g, James Gibsona,h, Emma Holdera,h,
Lisa Jonesa,h, Ann Dohertya,h, Rebecca Colesc,h, Stephanie G. Sumner-Jonesc,h, Marguerite Wasowiczd,h,
Michelle Manvelld,h, Uta Griesenbachd,h, Stephen C. Hydec,h, Deborah R. Gillc,h, Jane Daviesd,e,h,
D. David S. Collieb,h, Eric W.F.W. Altond,h, David J. Porteousa,h, A. Christopher Boyda,h
aMedical Genetics, School of Molecular and Clinical Medicine, University of Edinburgh, Western General Hospital, Edinburgh EH4 2XU, UK
bThe Roslin Institute and Royal (Dick) School of Veterinary Studies, Easter Bush Veterinary Centre, University of Edinburgh, Roslin EH25 9RG, UK
cGene Medicine Group, Nuffield Department of Clinical Laboratory Sciences, University of Oxford, John Radcliffe Hospital, Oxford OX3 9DU, UK
dDept of Gene Therapy, National Heart and Lung Institute, Imperial College, London SW3 6LR, UK
eDept of Paediatric Respiratory Medicine, Royal Brompton Hospital, London SW3 6NP, UK
fRockefeller University, New York, USA
gYale University, New Haven, CT 06519, USA
hUK Cystic Fibrosis Gene Therapy Consortium, United Kingdom
a r t i c l e i n f o
Article history:
Received 13 May 2009
Accepted 8 July 2009
Available online 15 July 2009
Keywords:
CFTR antibodies
Cystic fibrosis
Gene transfer
Nasal brushing cells
a b s t r a c t
Background: To assess gene therapy treatment for cystic fibrosis (CF) in clinical trials it is essential to
develop robust assays that can accurately detect transgene expression in human airway epithelial cells.
Our aim was to develop a reproducible immunocytochemical assay for human CFTR protein which can
measure both endogenous CFTR levels and augmented CFTR expression after gene delivery.
Methods: We characterised an antibody (G449) which satisfied the criteria for use in clinical trials.
We optimised our immunocytochemistry method and identified G449 dilutions at which endogenous
CFTR levels were negligible in CF samples, thus enhancing detection of transgenic CFTR protein. After
developing a transfection technique for brushed human nasal epithelial cells, we transfected non-CF and
CF cells with a clinically relevant CpG-free plasmid encoding human CFTR.
Results: The optimised immunocytochemistry method gave improved discrimination between CF and
non-CF samples. Transfection of a CFTR expression vector into primary nasal epithelial cells resulted in
detectable RNA and protein expression. CFTR protein was present in 0.05–10% of non-CF cells and 0.02–
0.8% of CF cells.
Conclusion: We have developed a sensitive, clinically relevant immunocytochemical assay for CFTR
protein and have used it to detect transgene-expressed CFTR in transfected human primary airway
epithelial cells.
Crown Copyright ? 2009 Published by Elsevier Ltd. All rights reserved.
1. Introduction
The cystic fibrosis transmembrane regulator gene (CFTR)
encodes a cAMP-regulated chloride channel that is localised to the
apical membrane of many epithelial cell types [1,2]. CFTR is
expressed in several organs including the lungs, pancreas, liver,
reproductive tract and sinuses. As well as controlling chloride
transport, the CFTR protein regulates other ion channels and
protein secretions [3–5].
CFTR mutations cause cystic fibrosis (CF), a common autosomal
recessive disease with a complex spectrum of phenotypes [6]. Most
CF-associated morbidity and mortality results from progressive
lung disease [7]. The most common CFTR mutation causes a dele-
tion of phenylalanine 508 (DF508). Modifier genes and environ-
mental effects also influence disease severity [8,9].
The aim of CF gene therapy is to express functional CFTR protein
in the lung airway epithelium. Progress has been slow due to
difficulties in transfecting epithelial cells in the lung milieu, short
* Corresponding author. Medical Genetics, School of Molecular and Clinical
Medicine,UniversityofEdinburgh,WesternGeneralHospital,EdinburghEH42XU,UK.
E-mail addresses: H.Davidson@ed.ac.uk,hdavids1@staffmail.ed.ac.uk(H.Davidson).
Contents lists available at ScienceDirect
Molecular and Cellular Probes
journal homepage: www.elsevier.com/locate/ymcpr
0890-8508/$ – see front matter Crown Copyright ? 2009 Published by Elsevier Ltd. All rights reserved.
doi:10.1016/j.mcp.2009.07.001
Molecular and Cellular Probes 23 (2009) 272–280

Page 2

duration of gene expression and a lack of effective assays to
measure gene delivery [10–17]. Despite these setbacks, data from
CF transgenic mice and human CF patients with milder phenotypes
suggest that 5–8% of normal CFTR mRNA levels can reduce lung
pathology [18,19]. Gene therapy therefore remains a realistic
option.
Levels of correction in initial trials of CF gene therapy are
expected to be low. Consequently a portfolio of sensitive assays will
be required for CFTR mRNA, protein and function before and after
gene transfer. Experiments on cultured cells, tissue samples and
primary nasal epithelial cells have given conflicting evidence about
levels and cellular localisation of the DF508 mutant protein
[20–26]. In cultured cells that experimentally over-express
DF508CFTR, the mutant protein is misfolded and retained in the
endoplasmic reticulum [20]. Studies comparing CFTR levels in
tissue samples from DF508 and non-CF individuals gave conflicting
results, with undetectable expression in DF508 sweat glands but
equal expression in DF508 and non-CF respiratory and intestinal
tracts [21–23,27–29]. A recent study using newantibodies found no
apically localised signal in the lung tissue of DF508 homozygous
individuals [24]. However, analysis of CFTR expression in brushed
DF508 nasal epithelial cells confirmed an apical signal, albeit at
a reduced level compared to non-CF samples [25,26]. Since DF508
individuals may have variable levels of apical CFTR protein in
airway epithelial cells, the design of a reliable assay to detect CFTR
before and after gene transfer is of paramount importance for the
development of effective genetherapy protocols. Anyassay must be
able to detect variable endogenous CFTR levels in DF508 patients as
well as augmented signal from the expressed transgene.
Here we describe progress towards developing an immunocy-
tochemical assay which gives improved sensitivity in detecting
endogenous apical CFTR in normal and CF nasal epithelial cells.
Using a new plasmid that directs sustained expression of hCFTR
mRNA in the mouse lung [30], we successfully transfected CF nasal
epithelial cells. Our studies provide valuable data totake forward to
clinical trials of CF gene therapy.
2. Materials and methods
2.1. Nasal and bronchial brushings
Airway cells were obtained with informed consent from
controls and CF patients using protocols approved by Lothian
Research Ethics Committee (nasal brushings) and Royal Brompton,
Harefield and National Heart and Lung Institute Ethics Committee
(bronchial brushings). Patients were screened using the standard
test for the 31 most common CFTR mutations. To accurately reflect
the genotypic diversity of patients in our forthcoming clinical trial,
we did not restrict the present study to subjects of a particular
genotype. For culture, nasal brushings [31] were collected in 2 ml
warmed BEGM (Lonza, Tewkesbury, UK) containing 1000 u/ml
penicillin and streptomycin then transferred to collagen-coated
96-well plates. Cells were harvested, fixed and adhered to silane
glass slides by cytospin [26]. For CFTR depletion, freshly brushed
nasal cells were incubated in 96-well plates with wiskostatin
(Calbiochem, San Diego, USA) diluted in 200 ml/well BEGM for 1 h
on ice prior to immunocytochemistry. Bronchial brushings were
taken by bronchoscope from non-CF and CF subjects. See
Supplementary Data for further details.
2.2. Coding and blinding
All procedures were carried out with sample identities known
to investigators apart from the immunocytochemical analyses
summarised in Fig. 6A which were conducted blind (see Supple-
mentary Data).
2.3. Plasmids and transfections
The CpG-free plasmids pG4-hCEFI-soLux, pG4-hCEFI-soCFTR2
and pG4-hCEFI-soEGFP express luciferase, human CFTR and EGFP
cDNAs respectively under control of the human CMV enhancer and
the elongation factor 1 alpha promoter [11,30]. The CFTRexpression
plasmid pCF-1-CFTR was constructed as described [32]. Plasmid
DNAs prepared using Endo-free columns (Qiagen, Crawley, UK)
were complexed with lipofectamine 2000 (Invitrogen, Glasgow,
UK) following the manufacturer’s protocol. Primary human nasal
epithelial cells (CF and non-CF) were grown to 75% confluence
(2–8 d). Cells that were to be transfected then received 0.2 mg
plasmid DNAcomplexed with 0.5ml lipofectamine; non-transfected
wells received lipofectamine alone. Cells were harvested 1 d post-
transfection for analysis of RNA (see below) or protein. For the
luciferase plasmid, sample medium was replaced with 100 ml
Reporter Lysis Buffer (Promega, Madison, WI, USA) then processed
for luciferase assay [31]. For the EGFP plasmid, transfection levels
were scored by examining w500 cells in at least 5 fields of view.
For the CFTR plasmid, immunocytochemistry was performed
(see below).
A 20 ml dose of pCF-1-CFTR plasmid DNA complexed with GL67
lipid (GL-67/DOPE/DMPE-PEG5000) [32] was administered to
anaethesised sheep maintained in a negative pressure ventilation
system [33] using a PARI LC plus jet nebuliser. At necroscopy, lung
segments wereinflated with 2:130% sucrose:OCT frozenand stored
at ?70
performed as described for nasal brushings [31].
?C. 8mm sections werecut, fixed and immunohistochemistry
2.4. Antibodies and immunocytochemistry (ICC)
Antibodies were diluted in 0.5% (w/v) BSA/PBS. CFTR antibody
G449 (0.68 mg/ml) was used at working dilution (1:100) unless
otherwise stated. Each batch was tested by Western blotting on
Calu3 cells and on the purified CFTR peptide against which it was
raised [31] (R domain, CFTR amino acids 653–716) (Supplementary
Fig. S1). Antibody MATG1061 (NBD1, CFTR amino acids 503–507/
509–515, 9 mg/ml; RD Biotech, Besançon, France) was used at 1:100
[31]. Control antibodies are described in Supplementary Data.
Secondary antibodies were conjugated to Alexa Fluor 594 and 488
(Molecular Probes, Invitrogen) and diluted to 1:800.
Data in Fig. 3A and B were obtained using our published ICC
protocol [31]. Subsequently we used a modified protocol (see
Results and Supplementary Data). All experiments included
a control slide without primary antibody. About 500 cells were
scored for each sample. Immunofluorescence was observed using
an Axioskop fluorescence microscope (Zeiss, Jena, Germany) with
a Coolsnap HQ camera (Photometrics, Tucson, AZ, USA). Images
were captured using SmartCapture2 software (Digital Scientific,
Cambridge, UK).
2.5. RNA preparation and TaqMan assays
Samples transfected with pG4-hCEFI-soCFTR2 and pG4-hCEFI-
soLux were stored in RNA Later buffer (Qiagen, Crawley, UK) before
RNA extraction with RNeasy reagents (Qiagen) (Supplementary
Data). Mean RNA Integrity Numbers and concentrations were
determined using a 2100 Bioanalyser RNA 6000 Nano Labchip assay
(Agilent Technologies, Palo Alto, CA) prior to in-solution DNase
treatment (Ambion, Applied Biosystems, Warrington, Cheshire, UK)
and RNeasy clean-up. mRNA levels were determined by two-step,
real-time TaqMan RT-PCR using plasmid RNA-specific primers and
H. Davidson et al. / Molecular and Cellular Probes 23 (2009) 272–280
273

Page 3

fluorogenic probes ([30,34] and Supplementary Data) on an ABI
PRISM 7700 Sequence Detection System with Sequence Detection
System software v1.6.3 (Applied Biosystems). Results are reported
as 40-Ct (total TaqMan run of 40 cycles).
2.6. Signal intensity analysis
Original SmartCapture images were converted to greyscale
using ImageJ software (http://rsb.info.nih.gov/ij) and the mean
pixel intensity of each cell was calculated (Supplementary Data).
2.7. Statistical analyses
Data were assessed for normal distribution using the Kolmo-
gorov–Smirnov test. Parametric (unpaired t-test, Student’s two-
tailed t-test) and non-parametric (Mann–Whitney test, Wilcoxon
signed rank test) tests were carried out as appropriate using Prism4
software (see figure legends and Supplementary Data). The null
hypothesis was rejected at p < 0.05. Power calculations were
carried out using 80% power based on the means of samples
(nQuery Advisor 6.01).
3. Results
3.1. Verification of CFTR antibody G449
Previously we tested 11 CFTR antibodies in human and ovine
nasal epithelial cells [31]. Only the polyclonal G449 satisfied our
three criteria for use in a clinical trial: availability in large quanti-
ties, reliability in quality control tests (detecting both the peptide
against which it was raised and CFTR protein in Calu3 cells), and
reproducibilityinimmunocytochemistry
(Supplementary Fig. S1 and Fig.1). G449 consistently detects apical
CFTR signal in human nasal epithelial (HNE) cells, as judged by
double-labelling withthe anti-CFTR
(Fig. 1K) and antibodies against cytokeratin, occludin, tubulin and
mucin (Fig. 1A, B, H, E and F respectively). Subsequently we
compared G449 antibody to the more recently published 528
antibody [24] but found that using our protocol, it did not meet our
criteria (Supplementary Fig. S2).
In some cells however, G449 also gave a nuclear signal (Fig.1A, J
and L). To characterise this phenomenon further we double-
labelled non-CF and CF nasal cells with G449 (raised to the R
domain of CFTR) and MATG1061 (raised to the NBD1 domain of
CFTR). The proportion of epithelial cells with apical signal was
similar with both (Table 1) and was lower in CF than non-CF
samples, as observed with G449 alone. Increasing concentrations of
wiskostatin, which disrupts actin and lowers the steady-state
surface CFTR pool [35], greatly reduced the percentage of cells with
apical G449 signal (Fig. 2). We concluded that G449 specifically
detects CFTR at the apical membrane and that the sporadic nuclear
staining is non-specific.
(ICC) experiments
monoclonalMATG1061
3.2. Immunocytochemistry: original methodology
We examined CFTR protein distribution in airwayepithelial cells
by ICC in 16 non-CF and 22 CF nasal brushing samples and 15 non-
CFand 7 CF bronchial brushing samples. Cells were double-labelled
with anti-cytokeratin antibody at 1:2000 and G449 at 1:100 or
1:600 (Fig. 3A and B) The 1:600 dilution was used to try to maxi-
mise discrimination between the two groups [25,26,31]. The
proportion of epithelial cells (cytokeratin-positive) expressing
apical CFTR was determined for each sample. Samples were also
analysed for epithelial, neutrophil, macrophage and other cell
content (Supplementary Results and Fig. S3).
Apical CFTR signal was detected in non-CF and CF epithelial
cells. Whilst, the proportion of positive cells was significantly
higher in non-CF than CF samples for both antibody dilutions, the
wide range of expression levels in both groups led to substantial
overlap between the groups. Discrimination between groups was
not improved by varying the primary antibody dilution.
Since mRNA levels of 5% may yield therapeutic results [18,19],
we performed a power calculation on the data in Fig. 3A and B
which showed that analysis of 4396 nasal samples and 1038
bronchial samples (1:100 antibody dilution) or 5292 nasal samples
(1:600 antibody dilution) would be required to detect a significant
increase of 5% in the proportion of CF cells with apical CFTR signal.
Therefore the existing ICC protocol was judged inadequate for
detecting modest increases in CFTR levels.
3.3. Immunocytochemistry: improved methodology
In a series of pilot experiments, we modified the ICC protocol by
increasingthe BSAconcentration in the blocking stepfrom1% to3%,
doubling the number of washing steps and diluting the cytokeratin
antibody to 1:10 000. We also used the Sequenza?immunostaining
system which ensures even application of antibodies by surface
tension. We compared the original and modified methods in four
CF and four non-CF samples using G449 dilutions of 1:100, 1:250
and 1:500. Within each sample the percentage of cells with apical
signal should decrease with increasing dilution of G449 antibody.
This decrease was observed consistently with the modified method
but was erratic with the original method. With the original method,
some lower antibody concentrations gave higher levels of apical
signal than higher antibody concentrations (arrows in Supple-
mentary Fig. S4B). The modified protocol gave lower background,
was more consistent and yielded a higher percentage of cells with
apical signal (Supplementary Fig. S4C).
Using the new ICC protocol, we extended the range of G449
dilutions to determine conditions under which endogenous CFTR
signal was negligible in CF samples, therebyoptimising detection of
recombinant CFTR. When 9 non-CF and 10 CF samples were
assessed for apical CFTR signal using G449 antibody at 1:100,1:500
and 1:750 we found improved separation between the two groups
(Fig. 3C). Endogenous CFTR signal was very low or negligible in CF
cells at G449 dilutions of 1:500 and above. Power calculations
showed that the minimum number of nasal brushing samples
required to demonstrate a significant increase of 5% in cells positive
for apical CFTR signal was 1806 for the 1:100 antibody dilution, 714
for 1:500 and 7 for 1:750. Therefore the improved ICC protocol,
together with greater antibody dilutions, provides discrimination
sufficient to detect modest increases in CFTR that might be
expected following gene therapy.
We also collated archive data using G449 antibody at 1:100
dilution from 61 controls and 28 CF patients (mixed genotype)
using the original ICC protocol and 68 controls and 31 CF patients
(mixed genotype) using the modified ICC protocol. Although all
samples were analysed during several different experiments,
together they constitute a valuable dataset because of the difficul-
ties in assembling a large collection of CF specimens. There was
a highly significant difference in the percentage of CFTR-expressing
HNE cells between the two groups (non-CF and CF) in both the
original and modified ICC protocol
p < 0.0001) despite considerable overlap (Fig. 4).
groups(bothgroups;
3.4. Transfection of non-CF human nasal epithelial cells
We next investigated whether the assay could detect plasmid-
derived CFTR protein against a background of endogenous CFTR
and simultaneously evaluate whether the clinical plasmid has the
H. Davidson et al. / Molecular and Cellular Probes 23 (2009) 272–280
274

Page 4

capacity to produce immunologically detectable CFTR in primary
human airway cells. We opted to transfect primary HNE cells and
chose lipofectamine 2000 as our transfection agentfor these invitro
studies since our experience has been that GL67 transfects very
poorly in vitro compared to in vivo. Cells were minimally handled
and allowed to settle on collagen-coated plates for 2–7 d before
transfection. Samples (n ¼ 4) were treated with the CpG-free
plasmid pG4-hCEFI-soCFTR2 (0.2 mg) complexed with lipofect-
amine or lipofectamine alone and harvested 1 d later for ICC with
antibody G449 at 1:250, 1:500 and 1:750.
During culture the cells gradually became non-polarised and
had varied morphology. The CFTR signal became cytoplasmic but
endogenous apical CFTR was apparent in some non-transfected
cells that had grown for 3 d or less (Fig. 5A). In transfected cells the
plasmid-derived CFTR signal was cytoplasmic and was too bright to
be captured using the same image exposure settings as were
applied in the case of endogenous CFTR (Fig. 5B–F). Epithelial cell
content exceeded 90% as judged by cytokeratin labelling of
untransfected control samples (n ¼ 3) harvested at the same time
as transfected cells.
Immunofluorescence from transfected cells was detectable at all
antibody dilutions; however endogenous CFTR was less bright at
1:500 and 1:750 (data not shown), allowing easier detection of
transfected protein. Transfection levels in the four samples, deter-
mined by counting epithelial cells with bright cytoplasmic signal,
were 0%, 0.25%, 2% and 0.5%. None of the untransfected cells gave
the bright signal characteristic of plasmid-expressed CFTR.
3.5. Transfection of CF nasal epithelial cells
Initially, nasal brushing samples from three CF subjects
(2 DF508/DF508, 1 DF508/G551D) and one non-CF control were
treated with 0.2 mg pG4-hCEFI-soCFTR2 (CFTR), 0.2 mg pG4-hCEFI-
soLux (luciferase), or no DNA (control) (n ¼ 2 wells per sample/
treatment). As well as being a negative control for CFTR detection in
ICC experiments, luciferase was included as a transfection and RNA
preparation control. ICC was carried out with antibody G449
diluted to 1:750.
Two of the CF samples failed to grow. The third sample (DF508/
DF508) gave vector-derived CFTR signal in 0.25–2% of cells which
Fig. 1. Endogenous CFTR expression in uncultured, untransfected human nasal epithelial cells. Nuclei are stained with DAPI (blue), although nuclei appear green, or have a green
tinge, in panels A, J and L because of a G449 labelling artefact (see text). All panels show non-CF human nasal cells apart from panel A which shows CF cells. (A) Endogenous apical
CFTR signal (arrow) in DF508/DF508 nasal epithelial cells double-labelled with G449 antibody (green) and cytokeratin antibody (red). (B) Endogenous apical CFTR signal (arrow) in
non-CF nasal epithelial cells double-labelled with G449 antibody (green) and cytokeratin antibody (red). (C) Absence of signal in negative control with secondary antibody but no
primary antibody. (D) Absence of signal with anti-b-galactosidase antibody (control). (E) Endogenous apical CFTR signal from G449 antibody (green) and cilia staining from a- and
b-tubulin antibodies (red, arrow). (F) Endogenous apical CFTR signal from G449 antibody (green) and secretory cell staining from mucin antibody (red, arrow). (G–I) Endogenous
apical CFTR signal from G449 antibody (green), tight junction staining from occludin antibody (red, arrow) and merged image of both antibodies. (J–L) Endogenous apical CFTR
signal from G449 antibody (green), anti-CFTR antibody MATG1061 (red, arrow) and merged image of both antibodies showing co-localisation.
H. Davidson et al. / Molecular and Cellular Probes 23 (2009) 272–280
275

Page 5

was comparable with 0.25–1.2% in the non-CF sample (data not
shown). All luciferase and no DNA controls were negative for
recombinant CFTR.
Three further experiments were conducted blind (with sample
identities hidden) on 6 CF and 11 non-CF samples. Each was treated
with pG4-hCEFI-soLux (luciferase), pG4-hCEFI-soCFTR2 (CFTR) and
no DNA (control) and assayed as before. When sample identities
were revealed, the luciferase and untransfected controls were
negative for recombinant CFTR. Of the CFTR-transfected samples,
4/6 CF and 8/11 non-CF samples were positive for recombinant
protein. Signal was observed in 0.05–10% of non-CF cells and
0.02–0.8% of CF cells. Fig. 6A represents data from wells containing
cells that reached >70% confluence (Figs. 5 and 6A).
TwotransfectedCFsamples(1DF508/G542X,11585-1G>A/P67L)
and three transfected non-CF samples were negative for plasmid-
expressed CFTR. These failed to reach the 75% confluence level
requiredforsuccessfultransfection.Overall,adequateconfluencewas
achieved in w70–90% of non-CF samples but only w40–60% of CF
samples, the lower rate resulting from contamination with bacterial
pathogens from the patient, reduced cell numbers obtained at
brushing and increased cell death during culture. Cell viability was
notmeasuredaswellscontainingcellswhichwereballedup,floating
and non-adherent to the collagen-coated wells were easily identifi-
able as of poor viability. However, if CF samples reached 70–90%
confluence, transfection levels were similar to those in non-CF
samples (Fig. 6A and B).
Three out of five pG4-hCEFI-soCFTR2-transfected non-CF
samples were positive for both vector-derived RNA and protein
(Table 2), demonstrating that the ICC assay detects plasmid-derived
CFTR protein. The no DNA controls (n ¼ 7) were all negative for
vector-derived CFTR protein, mRNA and luciferase (data not
shown). For pG4-hCEFI-soLux, 5/12 transfected samples were
positive for luciferase RNA and protein (Table 2) although some
protein-positive samples gave no detectable RNA. No RNA data are
available for CF samples because there were insufficient cells for
RNA extraction.
3.6. Transfection of ovine lung sections
As human ALIs are difficult to obtain and transfect and GL67
does not transfect cells well in vitro, we performed immunohisto-
chemistry (IHC) on ovine lung sections from sheep aerosolised with
lipid GL67 complexed with plasmid pCF-1-CFTR [32].
We have shown previously that the G449 CFTR antibody is
hCFTR-specific in ovine airways and detects protein expressed from
a similar hCFTR expression construct pCIKCFTR complexed with PEI
[31]. In the present study, the intensity of signal (Supplementary
Fig. S5) achieved using G449 on pCF-1-CFTR-transfected ovine lung
segments allowed us to discriminate transfected bronchioles
(Fig. 5G and H) from untransfected bronchioles (Fig. 5I) in a blinded
experiment. A sheep section with no primary antibody was
negative (data not shown).
3.7. G449 signal intensity in transfected HNE cells
ImageJ software was used to measure the difference in G449
signal intensity between putative transfected and non-transfected
HNE cells. 10 non-CF and 10 CF images at 40? magnification were
taken from samples 26 and 39 (non-CF) and 84 and 93 (DF508/
DF508) in the second blinded experiment. The mean pixel intensity
of the whole cell except the nucleus was calculated from a CFTR-
positive cell and a CFTR-negative cell within each image.
CFTR-positive cells had significantly greater signal intensity than
negative cells (8-fold greater for non-CF cells and 7-fold greater for
CF cells; both p < 0.0001) (Fig. 6B). ImageJ software was used to
measure the difference in G449 signal intensity between putative
transfected and non-transfected ovine epithelial lung cells in
a bronchiole (Supplementary Fig. S5). CFTR-positive cells had
significantly greater signal intensity than negative cells (4-fold
greater than non-transfected cells; p < 0.0001).
4. Discussion and conclusions
We have developed an immunocytochemical assay for detecting
CFTR protein following gene transfer. Methodological improve-
ments and extension of the range of antibody dilutions enabled us
to define conditions under which endogenous CFTR signal was low
or negligible, thus enhancing detection of plasmid-derived CFTR.
For the current batch of G449 antibody, the optimum dilution is
1:750. Each new batch of antibody would need to be titrated to find
the dilution at which discrimination between non-CF and CF apical
signal occurs. Power calculations on the data obtained with the
Table 1
Verification of CFTR antibody specificity G449 by double-labelling.
Subject and
genotype
No. of columnar
cells in sample
AntibodyColumnar cells
with apical signal
No.%
18
Non-CF
1805 G449
MATG1061
825
1013
45.7
56.0
19
Non-CF
530 G449
MATG1061
215
228
40.5
43.0
20
Non-CF
647G449
MATG1061
303
338
46.8
52.0
74
DF508/DF508
860G449
MATG1061
768.8
13.0112
75
DF508/DF508
1611 G449
MATG1061
327
203
20.2
12.6
76
DF508/DF508
1050
210
G449
MATG1061
251
38
23.9
18.1
The table shows the number of columnar epithelial cells in three non-CF and three
CF nasal brushing samples. The number and percentage of columnar cells with
apical signal after labelling with two CFTR antibodies, G449 and MATG1061, is also
shown. Samples 18, 19, 20, 74 and 75 were double-labelled with both CFTR anti-
bodies. Sample 76 was single-labelled with each antibody separately.
Fig. 2. Effect of wiskostatin on apical CFTR levels in nasal epithelial cells. Freshly
brushed nasal cells from 3 non-CF individuals (28, 37 and 39) were incubated with 0,
25, 50, 75 or 100 mM wiskostatin then fixed and stained with anti-CFTR antibody G449
at 1:100 dilution and anti-cytokeratin antibody at 1:5000 dilution. The mean
percentage of epithelial cells with apical CFTR signal was plotted. Bars indicate the
standard error of the mean.
H. Davidson et al. / Molecular and Cellular Probes 23 (2009) 272–280
276