Glycosylation differences between a cystic fibrosis and
rescued airway cell line are not CFTR dependent
Xiaosui
Jiang1,
Warren G.
Hill2,
Joseph M.
Pilewski2,3, and
Ora A.
Weisz1,2
1 Renal-Electrolyte Division
Laboratory of Epithelial Cell Biology,
3 Pulmonary and Critical Care
Division, Department of Medicine, and
2 Department of Cell Biology and
Physiology, University of Pittsburgh, Pittsburgh, Pennsylvania
15213
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ABSTRACT |
Altered
glycosylation of mucus and membrane glycoconjugates could explain
reported differences in binding of bacterial pathogens to cystic
fibrosis (CF) versus normal tissue. However, because bacteria can alter
cell surface glycoconjugates, it is not possible to assess the role of
cystic fibrosis transmembrane conductance regulators (CFTR) in
glycosylation in these studies. To address this issue, we have
developed quantitative lectin binding assays to compare cell surface
glycosylation in well-matched immortalized CF cells and rescued cell
lines. The CF airway bronchial epithelial cell line IB3-1 consistently
bound more peanut agglutinin (PNA) than its clonal derivative S9, which
stably expresses functional wild-type CFTR. Pretreatment with
neuraminidase increased PNA binding and abolished the difference
between the two cell lines. However, infection of the IB3-1 cells with
a replication-deficient recombinant adenovirus encoding CFTR restored
CFTR function but did not alter PNA binding to cells. In contrast,
treatment with the weak base ammonium chloride increased PNA binding to
both cell lines as expected. Our data show that even clonally related CF and rescued cells can exhibit significant differences in
carbohydrate processing. Although the differences that we found are
consistent with the proposed role for CFTR in modulating
intraorganellar pH, our data strongly suggest that they are CFTR
independent. These studies add a cautionary note to the interpretation
of differences in glycosylation between CF and normal primary tissues
and immortalized cells.
enzyme-linked lectin assay; sialylation; Golgi pH; Pseudomonas aeruginosa; cystic
fibrosis transmembrane conductance regulator
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INTRODUCTION |
THE PLEIOTROPIC EFFECTS of cystic fibrosis (CF) appear
to result from the mislocalization or impaired activity of an apical membrane chloride channel of epithelial cells, the cystic fibrosis transmembrane conductance regulator (CFTR). CFTR may also modulate intracellular chloride conductances and thus affect the pH of organelles along the secretory and endocytic pathways (4-6). An
elevated Golgi pH in CF cells could explain the observations that the
glycosylation profiles of proteins, glycolipids, and mucins differ
between CF and normal cells. In particular, it has been suggested that
CF glycoconjugates are more heavily sulfated and fucosylated than
normal and contain less sialic acid (6, 9, 17, 36). Because the pH
optimum for sialyltransferase is considerably lower than that of
fucosyltransferase or sulfotransferase [pH 5.8 vs. pH 6.8 for
sulfotransferase and pH 7.0-8.5 for fucosyltransferase (7, 27,
35)], these observed differences are consistent with the
predicted effects of increased intra-Golgi pH in CF cells. Furthermore,
because many strains of bacteria have been shown to bind avidly to
asialoconjugates, such as the glycolipid asialoGM1 (23), differences in
sialylation between CF and normal cells may explain the observation
that increased levels of bacteria bind to CF tissue and cells compared
with normal cells (22, 31, 32). However, it should be noted that, in
other studies, no difference in bacterial binding to cultures of CF and
non-CF nasal polyps was observed (8, 12).
Several issues complicate the studies of glycosylation and bacterial
binding in primary tissues. In general, it has been difficult to obtain
large enough quantities of normal mucin for detailed structural and
compositional studies (24, 29). Furthermore, not all of the
mucus-secreting cells in the airway express CFTR (16). Another problem
is that CF patients frequently have severe airway infection and
inflammation. Inflammatory mediators and bacteria have been shown to
enzymatically alter the composition of cell surface glycoconjugates and
mucous secretions, as well as the amount of mucin secreted (10, 21, 25,
29, 31, 32). Although these observations support the idea that the
surface glycosylation profiles of cells from CF patients differs from normal, the basis for these differences is not clear.
To avoid the problems of inflammation and bacterial contamination, we
have used a quantitative lectin binding assay to compare cell surface
glycosylation profiles of CF and rescued immortalized cell lines. For
our studies, we chose the CF bronchial epithelial cell line IB3-1 and
its clonally derived counterpart S9, which expresses functional CFTR.
Although we measured significant and reproducible differences in
terminal glycosylation profiles of these two closely matched cell
lines, our data suggest that these differences are independent of the
CF genotype of the cells.
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MATERIALS AND METHODS |
Cell lines. IB3-1 cells, derived from
CF bronchial epithelia and immortalized using adeno-12-SV-40 (38), and
S9 cells, generated from IB3-1 by transfection with a recombinant
adeno-associated viral vector encoding full-length, wild-type CFTR
(15), were kindly provided by Dr. Pamela Zeitlin. Cells were maintained
in F-12 medium supplemented with 10% fetal bovine serum and
penicillin/streptomycin.
Detection of functional CFTR by
6-methoxy-N-(3-sulfopropyl)quinolinium. Adenosine
3',5'-cyclic monophosphate (cAMP)-dependent anion efflux
was monitored by
6-methoxy-N-(3-sulfopropyl)quinolinium (SPQ; Molecular Probes, Eugene, OR) fluorescence changes in living cells (37). Briefly, subconfluent cells grown on 25-mm glass coverslips
were loaded with SPQ (10 mM) by 12 min of exposure at 37°C to
hypotonic NaI buffer (1:1 with water). Buffer composition (in mM) was
130 NaI, 4 KNO3, 1 Mg(NO3)2,
1 Ca(NO3)2,
10 glucose, and 20 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic
acid (HEPES), pH 7.4. Cells were mounted in a perfusion chamber placed
in a heating stage set to 37°C and were perfused with buffers
throughout the experiment. Imaging was performed on a Nikon Diaphot 300 inverted microscope equipped with a ×40 oil immersion objective
(Nikon CF fluor), image intensifier, and video camera. Excitation was at 330 nm, and image acquisition and analysis were performed by Metafluor software (Universal Imaging, West Chester, PA). The average
fluorescence intensity of individual cells in a field was monitored
every 15 s (8 frame average) throughout the assay, which ran for 14 min. Cells were perfused with isotonic NaI buffer for 2 min, nitrate
buffer (NaNO3 replaced NaI) for 4 min to assess the rate of iodide leakage/exchange from unstimulated
cells, 4 min in nitrate buffer supplemented with 10 µM forskolin and
200 µM 3-isobutyl-1-methylxanthine (IBMX), and then 4 min in iodide buffer to requench intracellular SPQ. Functional CFTR was detected as
an increase in the rate of dequenching of SPQ upon addition of
forskolin-IBMX. Assays on adenoviral-infected cells were performed using a blinded technique.
Enzyme-linked lectin assay. IB3-1 and
S9 cells were seeded in 96-well plates (6 wells/experimental group) and
were incubated at 37°C for 4 h in media. The cells were washed two
times with phosphate-buffered saline (PBS) containing 1 mM
CaCl2 and 1 mM MgCl2 at room temperature. Cells
in some wells were lysed with 1% Triton X-100 in PBS for determination
of total protein using the bicinchoninic acid assay (Pierce, Rockford,
IL). The remaining cells were fixed with 3% paraformaldehyde in PBS
for 15 min and then were washed with 0.2% gelatin (Sigma Chemical, St.
Louis, MO) in PBS containing 10 mM glycine (PBS-G). Endogenous
peroxidase activity was blocked before incubation with
lectin-horseradish peroxidase (HRP) conjugates by treatment with 0.3%
hydrogen peroxide in methanol for 15 min. The cells were then incubated
with 2 µg/ml of HRP-conjugated peanut agglutinin (HRP-PNA) or
HRP-conjugated wheat germ agglutinin (HRP-WGA; HRP-lectin conjugates
were from EY Laboratories, San Mateo, CA) in PBS-G for 30 min and then
washed with PBS-G. The bound lectin enzyme complex was visualized after light-protected incubation with 0.4 mg/ml of substrate
o-phenylenediamine (Sigma) and
hydrogen peroxide. The reaction was stopped by addition of 1 M
H2SO4,
and plates were read at 490 nm optical density using a microtiter plate
spectrophotometer (Molecular Devices, Sunnyvale, CA). In some
experiments, cells were pretreated with 0.05 U/ml of neuraminidase
(Calbiochem, La Jolla, CA) at 37°C for 30 min before fixation.
Adenoviral infection. IB3-1 cells were
mock infected or infected with replication-defective adenoviruses
encoding either nuclear-localized
-galactosidase (Ad-
-Gal) or
wild-type CFTR (Ad-CFTR) driven by the cytomegalovirus (CMV) promoter
[both viruses provided by Genzyme, Cambridge, MA (2, 28)]
at a multiplicity of infection (MOI) of 25-50. Two days
postinfection, cells were trypsinized and seeded into 96-well plates.
PNA binding was measured using 2 µg/ml HRP-PNA as described above.
The percentage of cells infected was monitored by histochemical
staining of
-Gal in cells infected with Ad-
-Gal as described
previously (26).
Immunoprecipitation and
phosphorylation. IB3-1 cells (uninfected or
infected with Ad-CFTR) and S9 cells were solubilized in lysis buffer
(1% Nonidet P-40 and 1 mM EDTA in 20 mM HEPES, pH 7) containing
protease inhibitor cocktail (10 mM leupeptin, 1 mM pepstatin A, 2 mg/ml
soybean trypsin inhibitor, 2 mg/ml aprotinin, 40 mg/ml
phenylmethylsulfonyl fluoride, and 0.2 mM dithiothreitol) for 1 min at
4°C. The cell lysates were precleared by incubation with normal
rabbit serum at 1:20 followed by precipitation using 25 µl of protein
A-Sepharose 6MB (Pharmacia Biotechnologies, Piscataway, NJ). After
centrifugation, supernatants were adjusted to 1× RIPA buffer
[1% Triton X-100, 1% sodium deoxycholate, and 0.1% sodium dodecyl sulfate (SDS) in 50 mM tris(hydroxymethyl)aminomethane (Tris) · HCl, pH 7.5]. The samples were
incubated with monoclonal antibodies against the COOH terminus and R
domain of CFTR (Genzyme) for 90 min, and antibody-antigen complexes
were isolated using protein A-Sepharose. After being washed
sequentially with RIPA buffer followed by 50 mM
Tris · HCl, pH 7.5, the Sepharose was resuspended in
protein kinase A (PKA) buffer (10 mM
MgCl2 in Tris buffer) and was
incubated with 5 units of cAMP-dependent PKA catalytic subunit
(Promega, Madison, WI) and 10 µCi
[
-32P]ATP (NEN,
Boston, MA) at 30°C for 60 min. The immune complex was released
from Sepharose by incubating in electrophoresis sample buffer
[125 mM Tris · HCl, pH 6.8, 5% (wt/vol)
SDS, and 25% (wt/vol) sucrose] containing 5% (vol/vol)
2-mercaptoethanol at 37°C for 5 min. The samples were run on a 6%
SDS-polyacrylamide gel electrophoresis (PAGE), and the dried gel was
exposed on X-ray film (X-OMAT AR; Eastman Kodak, Rochester, NY).
Statistical analysis. Results were
analyzed using the SigmaStat statistics program (Jandel Scientific, San
Rafael, CA). Data derived from three or more groups were compared by
one-way analysis of variance. Groups that differed significantly from
the control were identified using Dunn's test. Individual means from
three or more experiments were compared using the paired
t-test.
P < 0.05 was considered significant.
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RESULTS |
To determine the effect of CFTR on cell surface glycosylation, it
was necessary to confirm that the IB3-1 and S9 cells used in these
studies expressed the appropriate chloride secretory phenotype.
Functional CFTR activity was monitored using the halide-sensitive fluorophore SPQ as described in MATERIALS AND METHODS
(Fig. 1). S9 cells showed an increase in
the rate of SPQ dequenching upon stimulation with forskolin and IBMX,
indicating that these cells express functional CFTR (Fig.
1A). In contrast, cAMP stimulation cocktail had no effect on fluorescence in IB3-1 cells (Fig.
1B).

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Fig. 1.
cAMP-dependent anion permeability of IB3-1 and S9 measured using the
halide-sensitive fluorophore
6-methoxy-N-(3-sulfopropyl)quinolinium
(SPQ). Iodide efflux was measured in S9
(A) and IB3-1
(B) cells grown on glass coverslips
as described in MATERIALS AND METHODS. At time
(t) = 2 min, cells were switched to
nitrate buffer; at t = 6 min,
cAMP-stimulating cocktail was added; and at
t = 10 min, cells were returned to
iodide buffer. Arrows indicate the time at which buffers reached the
cells. S9 cells demonstrate an increased rate of SPQ dequenching upon
cAMP stimulation, suggesting that they express functional cystic
fibrosis transmembrane conductance regulator (CFTR), whereas the rate
of iodide leakage in stimulated IB3-1 cells is unchanged. Data are
normalized to 100% at the point of maximum dequenching and to 0 at the
2-min point when buffers are switched to nitrate buffer. Tracings
represent means ± SE from 12 cells
(A) and 16 cells
(B). Four S9 cells did not respond
to forskolin (not shown), suggesting that a population of these cells
may not express wild-type CFTR.
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Enzyme-linked lectin assays have been widely used for detection and
quantitation of terminal cell surface saccharides (3, 18). Within the
linear range, this sensitive assay allows direct comparison of lectin
binding to two cell lines, although the total number of lectin binding
sites cannot be measured. To determine whether CFTR expression might
influence the level of terminal sialylation in cells as previously
suggested, we measured the binding of HRP-PNA to IB3-1 and S9 cells.
This lectin binds preferentially to galactosyl(
-1,3)
N-acetylgalactosamine
[Gal(
-1,3)GalNAc] linkages found on
O-linked oligosaccharides such as mucin-type oligosaccharides, as well
as to the glycolipid asialoGM1. IB3-1 and S9 cells were seeded in
96-well plates, and HRP lectin binding was measured as described in
MATERIALS AND METHODS. Lectin binding was linear both with
respect to cell number per well and to the concentration of lectin
added (Fig. 2). Interestingly, HRP-PNA binding to IB3-1 cells was consistently higher than to S9 cells, suggesting that IB3-1 cells had increased levels of terminal galactose residues, as would be expected if sialyltransferase activity was reduced. The results were identical whether normalized to cell number
per well or total protein per well, suggesting that the two cell lines
did not differ markedly in size (not shown). Inclusion of 50 mM
galactose during the HRP-PNA incubation step completely abrogated
binding to both cell lines, suggesting that the binding we observed was
due to PNA interaction with cell surface glycoconjugates (not shown).
Furthermore, this result was highly reproducible and was observed
regardless of the passage number of the cells used (between
passages 20 and 35).

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Fig. 2.
Enzyme-linked lectin quantitation of cell surface galactose residues on
subconfluent IB3-1 and S9 cells using peanut agglutinin (PNA). Cells
(1-3 × 104/well; 4 wells each/time point) were plated in 96-well dishes. After 4 h, cells
were rinsed, fixed, blocked, and incubated with peroxidase-conjugated
PNA (1 and 2 mg/ml). After they were washed, cells were incubated with
o-phenylenediamine and peroxide, and
the amount of lectin-enzyme complex bound to the cells was quantitated
using a microtiter plate reader. O.D., optical density. Mean ± SD
for each condition is plotted. We consistently find that IB3-1 cells
bind more PNA than S9 cells (P < 0.05).
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To confirm that the difference in HRP-PNA binding to IB3-1 and S9 cells
was not an artifact due to differences in cell size or shape, we
compared binding of HRP-WGA to these cells. This lectin binds to
N-acetylglucosamine residues
(GlcNAc) and has high affinity for the
di-GlcNAc-containing core of N-linked oligosaccharides. Because the addition of the N-linked oligosaccharide core occurs cotranslationally in the endoplasmic reticulum, the level of WGA binding to cells should be independent of CFTR expression. As shown in
Fig. 3A,
binding of HRP-PNA to S9 cells was on average 46 ± 13% of binding
compared with IB3-1 cells (considered as 100%). By contrast, HRP-WGA
bound equally to IB3-1 and S9 cells (Fig. 3B).

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Fig. 3.
IB3-1 and S9 cells express similar levels of horseradish peroxidase
(HRP) conjugated-wheat germ agglutinin (WGA) cell surface binding
sites. IB3-1 and S9 cells (2 × 104/well) were incubated with 2 mg/ml HRP-PNA (A) or HRP-WGA
(B) and developed in the substrate
buffer as described in MATERIALS AND METHODS. Absorbance
readings were normalized against total protein recovered per well, and
the IB3-1 values were normalized to 100%. Data from 4 (A) and 3 (B) separate experiments are
expressed as means ± SE. Analysis using the paired
t-test shows a significant difference
in the binding of HRP-PNA binding to IB3-1 and S9 cells but no
difference in HRP-WGA binding to the two cell lines.
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We next wanted to determine whether the difference in PNA binding to
IB3-1 and S9 cells reflected a difference in the level of exposed
terminal galactose residues at the cell surface. To do this, we
measured PNA binding to cells before and after pretreatment with
neuraminidase. PNA binding to untreated S9 cells was 58% compared with
IB3-1 (Fig.
4A) as
observed previously. Upon neuraminidase treatment, binding to both cell
lines increased dramatically (Fig. 4B), and no significant difference
between the two cell lines was detected. These data indicate that the
percentage of exposed galactose residues in IB3-1 cells is
significantly greater than in S9 and support the idea that sialylation
in CF cell lines is impaired.

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Fig. 4.
PNA lectin binding to neuraminidase-treated IB3-1 and S9 cells is
similar. Cells were plated, mock treated
(A) or treated with 0.05 U/ml
neuraminidase at 37°C for 30 min
(B), and then processed as described
in MATERIALS AND METHODS. Neuraminidase-treated samples
were developed in substrate buffer for 3 min. Untreated samples were
developed for 20 min. Difference in PNA binding to IB3-1 compared with
S9 cells is statistically significant [means ± SE from 3 experiments (P = 0.01)], whereas PNA binding to the neuraminidase-treated cell
lines is not (P = 0.07). Results shown
are from a representative experiment that was performed 3 times with
similar results.
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To determine whether the difference in HRP-PNA binding to IB3-1 and S9
cells was CFTR dependent, we asked whether heterologous expression of
wild-type CFTR in IB3-1 cells would reduce the number of exposed
galactose residues at the cell surface. For these studies, we infected
cells with a recombinant replication-defective adenovirus encoding
CFTR. We tested the efficiency of infection of these cells using a
similar adenovirus encoding
-Gal. Cells were infected with
Ad-
-Gal at an MOI of 25, and
-Gal activity was detected colorimetrically after 2 days. Whereas uninfected cells showed no
staining (not shown), >90% of the virally infected cells stained positively for
-Gal activity (Fig. 5). Therefore,
under these conditions, we were able to infect the majority of cells.

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Fig. 5.
Recombinant adenoviruses efficiently infect IB3-1 cells. IB3-1 cells
were mock infected or infected with nuclear-localized -galactosidase
(Ad- -Gal), a recombinant adenovirus encoding -Gal, at a
multiplicity of infection (MOI) of 25. At 2 days postinfection, cells
were stained with X-Gal to determine the infection efficiency. Greater
than 90% of the cells express -Gal (inverted phase photomicrograph,
magnification ×100).
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To confirm that infection with Ad-CFTR resulted in the production of
full-length functional protein, we immunoprecipitated protein from
uninfected and virally infected cells (Fig.
6A). Cells were
infected with Ad-CFTR at an MOI of 25 and were incubated for 2 days. Because CFTR expression is driven by the butyrate-inducible CMV promoter in this virus, we also tested the effect of overnight induction with 1 mM butyrate on CFTR production. Cells were lysed and
CFTR was immunoprecipitated using commercially available
monoclonal antibodies directed against the COOH terminus and regulatory
domain of the protein. CFTR was detected on SDS-PAGE gels after in
vitro phosphorylation with PKA. As shown in Fig.
6A, lane
1, no CFTR could be detected in uninfected IB3-1 cells.
Upon infection with Ad-CFTR, radiolabeled bands consistent with the
known electrophoretic mobilities of the core glycosylated (Fig.
6A, band
B, ~130 kDa) and mature (Fig.
6A, band
C, ~180 kDa) forms of CFTR were observed (Fig.
6A, lane
2). Induction of CFTR expression in Ad-CFTR-infected IB3-1 cells with sodium butyrate dramatically increased the amount of
material in both bands (Fig. 6A,
lane 3; for clarity, the exposure shown is four times shorter than the other lanes). S9 cells expressed a
low but detectable level of both forms (Fig.
6A, lane
4).
To confirm that the CFTR expressed in these cells was functional, we
measured cAMP-stimulated halide efflux in virally infected cells using
the SPQ assay (Fig. 6B). In contrast
to uninfected IB3-1 cells or cells infected with Ad-
-Gal, cells
infected with Ad-CFTR and induced with sodium butyrate demonstrated a
dramatic cAMP-dependent increase in the rate of halide efflux,
indicative of functional CFTR expression. Together, these data
demonstrate that we were able to express functional CFTR in a large
proportion of the cells.

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Fig. 6.
IB3-1 cells express functional CFTR after infection with recombinant
adenovirus. A: subconfluent dishes of
IB3-1 cells were infected with wild-type CFTR (Ad-CFTR) at an MOI of
25. Before further analysis at 2 days postinfection, 1 dish was induced
overnight with 1 mM sodium butyrate. CFTR proteins were
immunoprecipitated using monoclonal antibodies against the COOH
terminus and R domain of the protein. Antibody-antigen complexes were
collected using protein A-Sepharose and were phosphorylated with
protein kinase A catalytic subunit and
[ -32P]ATP. Proteins
were separated by SDS-polyacrylamide gel electrophoresis, and
radiolabeled bands were detected by exposure to X-ray film.
Lane 1: uninfected IB3-1 cells;
lane 2: IB3-1 cells infected with
Ad-CFTR; lane 3: butyrate-induced
Ad-CFTR-infected IB3-1 cells; lane 4:
S9 cells. Positions of expected migration of the core glycosylated form
(band B) and mature form of CFTR
(band C) are noted.
Lanes 1,
2, and
4: 16-h exposure;
lane 3: 4- h exposure.
B: IB3-1 cells grown on glass
coverslips were mock infected (IB3) or infected with Ad-CFTR
(IB3-Ad-CFTR) or Ad- -Gal (IB3-Ad- -Gal) at an MOI of 50 and
induced with sodium butyrate as described above. Functional CFTR
activity of uninfected and infected cells was monitored using the SPQ
assay as described in MATERIALS AND METHODS. Arrows
indicate the time at which buffers reached the cells. Tracings
represent means ± SE from 19, 37, and 14 cells (IB3,
IB3-Ad- -Gal, and IB3-Ad-CFTR, respectively) and are normalized to
100% at the point of maximum dequenching of IB3-Ad-CFTR and to 0% at
the 2-min point when buffers were switched to nitrate buffer. All of
the IB3-Ad-CFTR cells responded to forskolin, suggesting that they
expressed functional CFTR.
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HRP-PNA binding to IB3-1, S9, and virally infected IB3-1 cells was
determined using the enzyme-linked lectin assay. The results (Fig.
7) show that CFTR
expression has no effect on lectin binding to IB3-1 cells. HRP-PNA
binding to IB3-1 cells was unaffected by infection with Ad-CFTR
regardless of whether CFTR expression was induced with sodium butyrate.
Neither adenoviral infection with Ad-
-Gal nor sodium butyrate
treatment of uninfected IB3-1 cells altered PNA binding significantly.
In contrast, HRP-PNA binding to S9 cells was clearly different compared
with IB3-1 cells (P < 0.0001). Thus
it appears that the difference between HRP-PNA binding to IB3-1 and S9
cells is CFTR independent.

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Fig. 7.
Infection with Ad-CFTR does not alter PNA binding to IB3-1 cells. IB3-1
cells were mock infected or infected with Ad-CFTR or Ad- -Gal at an
MOI of 25-50. At 2 days postinfection, cells were reseeded into
96-well plates, and HRP-PNA binding was measured after 4 h. Some dishes
were treated with 1 mM sodium butyrate for 24 h before
reseeding. Results were normalized against HRP-PNA binding to
uninfected IB3-1 cells (considered as 100%) in each experiment and are
presented as means ± SE. 1: Uninfected IB3-1 cells treated with
sodium butyrate (n = 3); 2: IB3-1
cells infected with Ad-CFTR (n = 4);
3: IB3-1 cells infected with Ad-CFTR and induced with sodium butyrate
(n = 4); 4: IB3 cells infected with
Ad- -Gal (n = 4); and 5: S9 cells
(n = 9). Whereas HRP-PNA binding to S9
cells is statistically different from IB3
(P < 0.0001), there is no
statistical difference between induced and/or infected IB3-1
cells compared with control (P > 0.1).
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To address the possibility that the 2-day infection period was not long
enough to remodel cell surface glycoconjugates sufficiently for
measurement, we synthetically altered intra-Golgi pH by incubating cells for 2 days in 10 mM ammonium chloride and measured the effect on
HRP-PNA binding (Fig. 8). This treatment,
which elevates the pH of all acidic intracellular compartments (11),
was not toxic to cells over this time period. PNA binding to both cell
lines increased measurably in ammonium chloride-treated cells relative to control untreated cells, suggesting that this treatment interfered with sialylation as expected. Interestingly, the difference in PNA
binding was preserved between ammonium chloride-treated IB3-1 and S9
cells, perhaps suggesting that the difference in terminal glycosylation
between these two cell lines is pH independent.

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Fig. 8.
Ammonium chloride treatment increases HRP-PNA binding to IB3-1 and S9
cells. IB3-1 and S9 cells were incubated with (+) or without ( ) 10 mM ammonium chloride for 2 days before quantitation of HRP-PNA binding.
Even after pretreatment with ammonium chloride, PNA binding to IB3-1 is
significantly different from binding to S9 cells (means ± SD;
P < 0.05). This experiment was
performed 3 times with similar results.
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DISCUSSION |
We have compared the glycosylation profile of a matched CF and rescued
epithelial cell line using a quantitative enzyme-linked lectin assay.
This assay was linear with respect to cell number (or protein
concentration) and concentration of HRP-conjugated lectin added. Using
this assay, we demonstrated a significant and highly reproducible
difference in the amount of terminal galactose present on CF versus
normal cells. This lectin recognizes Gal(
-1,3)GalNAc linkages
preferentially and thus is most specific for O-linked oligosaccharides
and glycolipids, particularly asialoGM1. When cells were pretreated
with neuraminidase, PNA binding to both cell lines increased
dramatically, and no significant difference was observed in the level
of total galactose residues on IB3-1 compared with S9 cells.
Furthermore, binding of WGA lectin was identical between IB3-1 and S9
cells. Together, our results show that IB3-1 cells have an increased
level of terminal galactose residues compared with S9 cells but
otherwise are similar in surface glycoconjugate composition. These data
validate the use of the enzyme-linked lectin assay as a quantitative
method to compare cell surface glycosylation profiles between related
cell lines.
Because our results supported the hypothesis that CF cells may have
increased intraorganellar pH, we investigated whether restoring CFTR
function to the IB3-1 cells would also rescue the glycosylation
phenotype. Infection with recombinant, replication-defective adenovirus
encoding CFTR for 2 days resulted in >90% transduction efficiency.
We were able to qualitatively measure CFTR in the infected cells using
immunoprecipitation followed by in vitro phosphorylation. Furthermore,
the CFTR expressed in these cells was functional as assessed by the
halide-sensitive fluorophore SPQ. However, expression of CFTR did not
alter the glycosylation profile of IB3-1 cells, suggesting that the
decreased sialylation that we observed in this cell line was CFTR
independent.
Given the large difference in PNA binding that we observed between
IB3-1 and S9 cells and the estimated 90% adenovirus infection efficiency, we should have easily been able to detect a change in PNA
binding if CFTR expression altered oligosaccharide processing in IB3-1
cells. One argument against our interpretation is that glycosylation turnover is too slow to detect differences by 2 days
postinfection. Several studies have examined the half-life of membrane
proteins in other cultured cell lines (1, 14, 19). Although the results
of these studies suggest that the rate of protein turnover is somewhat
dependent on cell type and cell growth conditions, protein turnover in
rapidly dividing cells was generally found to follow diphasic kinetics,
with between 25 and 35% of protein turning over rapidly (half-life of
between 1 and 10 h) and the remainder being degraded more slowly
(half-time between 1 and 5 days). These measurements do not take into
account the synthesis of new membrane proteins required for continuing cell division and therefore represent an underestimate of the rate of
protein remodeling in dividing cells. Under our growth conditions,
IB3-1 cells divide approximately every 24 h, even after adenoviral
infection. Therefore, we estimate that at least 75% of the total
protein is newly synthesized within 48 h of infection. Because mRNA
from recombinant adenoviruses can be detected within 6 h of infection
(20), the majority of glycoconjugates synthesized in IB3-1 cells during
the 2 days of postinfection incubation will have been synthesized in
cells expressing wild-type CFTR. Furthermore, treatment with ammonium
chloride for 48 h increased PNA binding to cells by ~30%, thus
confirming that significant remodeling of the cell surface carbohydrate
profile occurred during this period. We attempted to extend the length
of time in culture after adenoviral infection; however, because the
cells continue to proliferate and the adenovirus-mediated gene transfer
is episomal (20), the percentage of transduced cells declined
dramatically with time in culture. After 4 days, only 55% of cells
infected with Ad-
-Gal showed detectable
-Gal staining. At this
time point, there was no difference in PNA binding between
Ad-CFTR-infected and uninfected IB3-1 cells (not shown). In addition,
we tried to express endogenous functional CFTR at the cell surface of
IB3-1 cells using other nonviral approaches, including incubation for 2 days with 0.5 mM 4-phenylbutyrate (30), incubation at reduced temperature (27°C; see Ref. 13), and treatment with 0.1 mg/ml geneticin, which can promote read through from the W1282X allele present in IB3-1 cells (M. Howard and M. Gondor, personal
communication). However, in our hands, none of these treatments
produced detectable functional CFTR at the cell surface, as monitored
by SPQ.
The increased level of terminal galactosylation in IB3-1 cells could
indicate an increased number of bacterial binding sites on these cells
compared with S9 cells. Interestingly, Imundo et al. (22) recently
reported that IB3-1 cells bind two times as many
Pseudomonas aeruginosa bacteria as
another subclone (C38) that expresses functional CFTR. This observation
may suggest that this subclone also expresses lower levels of
sialylated glycoconjugates than IB3-1 cells.
There are several possible reasons for the differences in terminal
galactosylation that we observed between IB3-1 and S9 cells. The most
likely explanation is that the glycosylation pattern of the clonally
derived S9 cells reflects the profile of a subpopulation of the
original IB3-1 population from which it was generated. IB3-1 cells are
polyploid and contain between 80 and 90 chromosomes. Thus any subclone
of this heterogeneous population may express different levels or
isoforms of any of the myriad cellular components that affect
glycosylation, such as glycosyltransferases or sugar transporters.
Because ammonium chloride did not equalize PNA binding between IB3-1
and S9 cells, we suspect that the difference in glycosylation between
the two cell lines is pH independent. The difference in glycosylation
is not likely to be due to differences in cell passage number, since we
observed no difference in glycosylation of either line with increased
passage and since the levels of WGA binding to the cells remained
identical. Furthermore, in a detailed study examining oligosaccharide
composition and structure, Swiedler et al. (33, 34) found virtually no
change in the glycosylation profile of cells even after up to 1 yr of
continuous passage.
Our data suggest that differences in glycosylation observed between CF
and control cell lines must be interpreted with caution, as such
differences are not necessarily CFTR dependent. To address the role of
CFTR in glycosylation, it will be necessary to compare CF and corrected
primary tissue from a single individual. The enzyme-linked lectin assay
described here appears to be sensitive and reproducible enough to be
used for such studies.
 |
ACKNOWLEDGEMENTS |
We thank Drs. Edward Wing and Steven Gregory for access to their
microtiter plate reader, Dr. Pamela Zeitlin for the IB3-1 and S9 cell
lines, and Drs. Sam Wadsworth and Donna Armentano at Genzyme for
providing replication-defective recombinant adenoviruses.
 |
FOOTNOTES |
This work was supported in part by grants from the Cystic Fibrosis
Foundation to O. A. Weisz and J. M. Pilewski.
Address for reprint requests: O. A. Weisz, Renal-Electrolyte Division,
University of Pittsburgh, 3550 Terrace St., Pittsburgh, PA 15213.
Received 30 May 1997; accepted in final form 18 July 1997.
 |
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