From the Programme in Cell Biology and Genetics at the Hospital for Sick Children and the Departments of Physiology and Molecular Genetics at the University of Toronto, Toronto, M5G 1X8 Ontario, Canada
Received for publication, July 27, 2000, and in revised form, November 26, 2000
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ABSTRACT |
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It has been previously determined that ClC-2, a
member of the ClC chloride channel superfamily, is expressed in certain
epithelial tissues. These findings fueled speculation that ClC-2 can
compensate for impaired chloride transport in epithelial tissues
affected by cystic fibrosis and lacking the cystic fibrosis
transmembrane conductance regulator. However, direct evidence linking
ClC-2 channel expression to epithelial chloride secretion was lacking. In the present studies, we show that ClC-2 transcripts and protein are
present endogenously in the Caco-2 cell line, a cell line that models
the human small intestine. Using an antisense strategy we show that
ClC-2 contributes to native chloride currents in Caco-2 cells measured
by patch clamp electrophysiology. Antisense ClC-2-transfected
monolayers of Caco-2 cells exhibited less chloride secretion (monitored
as iodide efflux) than did mock transfected monolayers, providing the
first direct molecular evidence that ClC-2 can contribute to chloride
secretion by the human intestinal epithelium. Further, examination of
ClC-2 localization by confocal microscopy revealed that ClC-2
contributes to secretion from a unique location in this epithelium,
from the apical aspect of the tight junction complex. Hence, these
studies provide the necessary rationale for considering ClC-2 as a
possible therapeutic target for diseases affecting intestinal chloride
secretion such as cystic fibrosis.
The physiological significance of ClC-2, a ubitquitously expressed
member of the ClC family of chloride channels (1) is not fully
understood. Based primarily on studies of native ClC-2 message and
protein expression, roles for ClC-2 in neuronal and epithelial tissue
have been proposed (2, 3). ClC-2 channel activity has been implicated
in the regulation of neuronal responses to GABA-A receptor interaction
(2). In non-neuronal cells, ClC-2 function has been linked to volume
regulation, and in epithelial cells, it has been linked specifically to
chloride secretion (3).
Immunolocalization studies of ClC-2 revealed that it is situated on the
apical surface of airway epithelial cells in neonatal rat airways (4).
Subsequent Ussing chamber studies showed that luminal acidity promoted
chloride secretion in neonatal airway cells via a cadmium-sensitive
channel (5). These findings prompted us to suggest that ClC-2, a
channel that exhibits these properties, may mediate chloride secretion
in neonatal rat airways. In addition, Schwiebert et al. (6)
reported that ClC-2-like currents are present in airway epithelial
cells derived from an adult patient with cystic fibrosis
(CF)1 and suggested chloride
transport via ClC-2 may be able to compensate for defective or absent
CFTR chloride channels in the CF airway epithelium. Further, because
ClC-2 message can be detected in intestinal epithelial tissue obtained
from cftr-knockout mice, Joo et al. (7) also
suggested that there is the potential for ClC-2 to provide a bypass
pathway for chloride transport in CF affected intestines. Despite these
intriguing observations, there was no direct molecular evidence to
suggest that ClC-2 contributes to native chloride secretion.
In the present work, we assessed the role of ClC-2 in chloride
secretion in the Caco-2 cell line, a cell line that models the human
small intestinal epithelium (8, 9). Using immunofluorescence and
confocal microscopy, we confirmed that ClC-2 protein is endogenously expressed in the plasma membrane of Caco-2 cells. Further, we show that
it is uniquely situated at the apical aspect of the tight junctions
between cells in fully differentiated monolayers of these cells. Using
an antisense strategy, we show that endogenously expressed ClC-2
mediates currents across the plasma membrane of single Caco-2 cells
and, finally, that ClC-2 can contribute to native anion secretion
across Caco-2 cell monolayers.
Caco-2 Cell Culture--
Caco-2 cells were obtained from the
American Type Culture Collection (Manassas, VA). They were grown in
Earl's Northern Analysis--
Total RNA was isolated from Caco-2 cell
monolayers using the Trizol method as recommended by the supplier (Life
Technologies, Inc.). RNA (2 µg) was analyzed on agarose gels (1%)
containing 0.6 M formaldehyde and transferred to Hybond-N
membranes (Amersham Pharmacia Biotech). Blots were cross-linked with UV
radiation and hybridized with mouse-specific ClC-2 cDNA fragments
radiolabeled by random priming (10). Final conditions of washing
included 0.2× SSC (sodium chloride/sodium citrate) with 0.1% SDS at
60 °C. The blots were exposed to X-Omat film (Kodak) for 24-72 h at
Western Analysis--
Caco-2 cells, grown on 60-mm plastic
dishes, were washed with phosphate-buffered saline containing 10 mM mono-dibasic mix, pH 6.8, and 150 mM NaCl,
final pH 7.2, and incubated with 900 µl of lysis buffer containing
1% Triton X-100, 120 mM NaCl, 10 mM Tris, 25 mM KCl, 25 mM MgCl2, 1.8 mM CaCl2, and protease inhibitors leupeptin (10 µg/ml) and apotinin (10 µg/ml), 1 mM benzamidine, 10 µM E64, and 2 mM dithiothreitol. The cells
were scraped off, and the suspension was mixed by vortexing. The
supernatant was then centrifuged for 10 min at 4 °C at 48,000 × g to isolate a crude membrane preparation. Following
protein assay of the supernatant, 50 µg of this preparation was
analyzed by SDS-polyacrylamide gel electrophoresis (8% gel) using
anti-ClC-2 antibody at a concentration of 2 µg/ml. This polyclonal
antibody, as described previously, was generated against a GST fusion
peptide containing amino acids 31-74 of rat ClC-2 (rClC-2 cDNA
kindly provided by T. Jentsch). The ClC-2-specific antibody was
immunopurified from a matrix of GST-N-peptide coupled on an activated
agarose column as previously described (11, 12). The monoclonal
antibody against Immunofluorescence--
Immunofluorescence labeling was
performed on Caco-2 cells grown on 35-mm circular coverslips or on
clear 35-mm, 0.4-µm pore Snapwell (Corning Costar) filters. The
pattern of ClC-2 labeling was identical regardless of the support
employed. Cells were fixed with paraformaldehyde AM (4% in
phosphate-buffered saline) and permeabilized with 0.5% Triton X-100 in
phosphate-buffered saline. Cells were incubated for 0.5 h at
25 °C in 5% normal goat serum, 0.05% Triton X-100 in Tris-buffered
saline containing 10 mM Tris-Cl and 150 mM NaCl
with final pH 8, and then for 2.5 h with the polyclonal antibody
against ClC-2 (30.7 µg/ml) or overnight in the refrigerator with the
polyclonal antibody against ClC-3 (30 µg/ml) (Alomone Labs Ltd.,
Jerusalem, Israel). Then the cells were washed and incubated with Texas
Red-conjugated or fluorescein isothiocyanate-conjugated anti-rabbit
secondary antibody (0.02 mg/ml; Molecular Probes) and washed again
before mounting. For colocalization studies of ClC-2 and the tight
junction protein occludin, the above procedure was followed by
additional incubation with the monoclonal anti-occludin antibody (0.002 mg/ml, Zymed Laboratories Inc., Missisauga, Canada) for 1 h and washed. Cells were then incubated with Texas
Red-conjugated anti-mouse secondary antibody (0.02 mg/ml; Molecular
Probes) and washed before mounting. For the competition studies, the
anti-ClC-2 antibody was preincubated with 2-fold excess of the
antigenic fusion peptide overnight at 4 °C before incubation. Slides
were viewed on an Olympus Vanox AHBT3 microscope using epifluorescence, and images were captured using the Image Pro Plus program (Cybernetics, L.P.). For confocal microscopy, sections (each 0.7 µm in thickness) were viewed with a 100x objective a Leica TCS 4D microsope, and the
images were captured using the SCANware 5.01 program.
Patch Clamp Studies of Caco-2 Cells--
Caco-2 cell membrane
currents were measured using conventional whole cell patch clamp
technique (13). Patch clamp electrodes were prepared from borosilicate
glass capillaries (outer diameter, 1.5 mm; inner diameter, 1.18 mm)
with an inner filament (World Precision Instruments, Inc.,
Sarasota, FL) on a Narishige PP-83 patch electrode puller using the
standard two-pull technique. The tip resistance was 3-5 M
ClC Constructs--
The ClC-2 sense construct was made by
directional cloning of the rat (r)ClC-2 open reading frame (kindly
provided by T. Jentsch, Hamburg, Germany) with BamHI (5')
and EcoRI (3') linkers into the BamHI and
EcoRI restriction sites of the eukaryotic vector pCDNA
3.1 (+) (Promega, Madison, WI). The rat ClC-2 sequence shares 77%
identity with the human sequence at the nucleotide level. The antisense
ClC-2 construct was made by cloning the ClC-2 open reading frame into
pCDNA 3.1( Intranuclear Injection of Plasmid--
Caco-2 cells were
microinjected with plasmids at day 1 after plating on glass coverslips
for patch clamp experiments. In this procedure, the Eppendorf
microinjector 5246 system, the micromanipulator 5171 system, and a
Nikon Diaphot inverted microscope were used. Nuclear microinjection was
performed with the Z (depth) limit option, using 0.3-s injection
duration and 40-60 hPa injection pressure. Injection femtotips were
pulled from borosilicate glass capillaries with an internal diameter of
0.5 ± 0.2 µm. Plasmids were diluted to a final concentration of
50 µg/ml for sense ClC-2 plasmids, 50 and 300 µg/ml for antisense
ClC-2, and 300 µg/ml antisense ClC-4. The injection buffer contained
in 50 mM HEPES, 50 mM NaOH, 40 mM
NaCl, pH 7.4. Fluorescein isothiocyanate-labeled dextran (0.5%, Sigma)
was also added to the injection medium to identify successfully
microinjected cells.
Transient Transfection of Caco-2 Cell Monolayers--
For
transfection of Caco-2 cells with antisense ClC-2 DNA, the Lipofectin
transfection protocol was followed (Life Technologies, Inc.). Briefly,
~2 × 105 cells were seeded on 35-mm tissue culture
plates in culture medium supplemented with serum. Cells were then
incubated at 37 °C in a 5% CO2 incubator overnight to
allow them to reach 80% confluency (~106 cells/plate).
Two solutions, one containing 2 µg of antisense ClC-2 cDNA
(dissolved in serum-free medium) in 100 µl of OPTI-MEM I reduced
serum medium and the second containing 20 µl of Lipofectin reagent in
100 µl of OPTI-MEM I reduced serum medium were mixed and incubated at
room temperature for 45 min. Following addition of 800 µl of OPTI-MEM
I reduced serum medium, this transfection mixture was applied to the
cells (after washing them with serum-free medium). The cells were then
incubated at 37 °C in a 5% CO2 incubator for 48 h,
after which the transfection medium was replaced with normal medium
containing 10% serum and antibiotics (as described above). 24 h
later, cells were harvested for immunoblot analysis or studied by
iodide efflux assay.
Ussing Chamber Analysis--
Short circuit current measurements
were performed on Caco-2 cells grown to confluency on Snapwell clear
filters, with a surface area of 1.13 cm2 (Corning Costar).
The average transepithelial resistance of the cells used was 2421 ± 357.2 Iodide Efflux--
Caco-2 cells grown on coverslips (80%) were
transfected either with antisense ClC-2 plasmid in the pCDNA vector
or with vector alone as described under "Experimental Procedures."
Only monolayers possessing 1 × 106 cells after the
entire transfection protocol were used for subsequent assays. The
transfected cells were iodide loaded according to established methods
(14, 15) using a 1-ml Ringers Nitrate loading buffer at pH 7.4 containing 136 mM NaI, 4 mM KNO3, 2 mM CaNO3·4H2O, 2 mM
MgNO3·6H2O, 11 mM glucose, and 20 mM HEPES. The cells were incubated for 1 h at 37 °C
in the presence of 5% CO2 in the above buffer. After this
incubation period, the coverslips covered with iodide-loaded cells were
washed for a total of 15 s in three separate baths containing
Ringers nitrate efflux buffer (110 mM NaNO3, 4 mM KNO3, 2 mM
CaNO3·4H2O, 2 mM
MgNO3·6H2O, 11 mM glucose, and 20 mM HEPES, pH 7.4) with the osmolarity adjusted to 300 mOsm
with added sucrose. After this washing period, efflux of cellular
iodide was assessed continuously after placing the coverslip into an
isotonic solution (the above Ringers Nitrate efflux buffer) or
hypotonic solutions (osmolarity adjusted 228 mOsm). Iodide efflux
(measured as a change in mV) was assessed over a 5-min period, using an
iodide sensing electrode (Fisher). Changes in voltage were acquired
using the FETCHEX data acquisition program (pCLAMP 6.04, Axon Inst.)
and data analyzed using FETCHAN software.
Statistics--
Patch clamp measurements are presented as the
means ± S.E. Most statistical analyses were performed using the
Student's unpaired test. Results obtained in Ussing chamber studies
and in Patch clamp studies with hypotonic shock were analyzed using the
Student paired test. Differences between two groups were considered
significant with p values <0.05.
ClC-2 Message and Protein Are Expressed in Caco-2 Cells--
ClC-2
mRNA in Caco-2 cells was detected by Northern blot analysis as a
4.6-kb transcript (Fig. 1A).
This size transcript plus a smaller transcript of ~3.3 kb in size has
been detected in several other tissues and cell lines, as well,
including the colonic epithelial cell line T84 (6).
Immunoblots (Fig. 1B) using a polyclonal antibody directed
against ClC-2 (12) showed that ClC-2, migrating as a 90-97-kDa
protein, is expressed in Caco-2 cells. This signal was competed using
the GST-ClC-2 fusion peptide against which the antibody was raised, not
GST alone, indicating its specificity for ClC-2.
Immunofluorescence labeling using the above antibody suggests that
ClC-2 protein localizes to plasma membrane and/or submembranous vesicles in Caco-2 cells (Fig. 2).
Further, this signal was specific because it was competed using the
antigenic peptide described above.
ClC-2 Is Functionally Expressed on the Plasma Membrane of Caco-2
Cells--
Whole cell patch clamp studies were performed to determine
whether ClC-2 is functional in the membrane of Caco-2 cells. Because previous studies in heterologous systems showed that ClC-2 expression conferred chloride currents were activated by hyperpolarization and by
hypotonic shock (1, 11, 16), we assessed whether these manipulations
could activate ClC-2 endogenously expressed in Caco-2 cell membranes by
patch clamp electrophysiology. We functionally isolated
anion-dependent currents by using intracellular (pipette)
and extracellular (bath) solutions which contained NMDG chloride as the
predominant salt. We chose a voltage step protocol that has been used
in previously published studies of ClC-2 (11, 16). Briefly, from a
holding potential of
As shown in Fig. 3 (B and C), whole cell chloride
currents in Caco-2 cell were stimulated by 25% hypotonic shock. We
detected an increase in membrane currents from
We used an antisense strategy to confirm that the above currents were
mediated by ClC-2 because the pharmacological approach lacks
specificity. First, we confirmed that transient transfection of ClC-2
antisense cDNA (see "Experimental Procedures") successfully reduced ClC-2 protein expression by Western analysis of cell lysates from ClC-2 antisense-transfected Caco-2 cells. Using the NIH Imaging Program, we found that there is a 70% decrease in ClC-2 protein quantity in antisense ClC-2 transfected Caco-2 cells relative to
control (vector-alone transfected cells). We verified by assessing
For patch clamp studies, we manipulated ClC-2 expression using
intranuclear plasmid injection technique (20, 21), because this method
permits control of plasmid copy number and hence has greater precision
in manipulating the level of antisense expression. Fluorescein
isothiocyanate-dextran was coinjected with the plasmid to permit
identification of manipulated cells. We found that microinjection of
antisense ClC-2 cDNA into Caco-2 cells decreased the ClC-2-like currents in a dose-dependent manner (Fig.
5, A and B). The
negative whole cell current measured at
Inhibition of hyperpolarization-activated chloride currents by ClC-2
antisense expression was a specific response, because nuclear injection
of antisense ClC-4, a distinct membrane of the ClC chloride channel
family (3), did not affect these endogenous currents (Fig.
5C). The currents measured in antisense ClC-4 injected Caco-2 cells (
To determine the contribution of ClC-2 to HTS-stimulated chloride
currents in Caco-2 cells, we studied the effects of antisense ClC-2
transfection on the development of hypotonicity-activated chloride
currents in Caco-2 cells. As shown in Fig. 5E, the amplitude of the HTS-stimulated chloride currents measured in noninjected Caco-2
cells at -160 mV (-65 ± 8 pA/pF, n = 4) was
almost four times larger than the currents measured in antisense ClC-2
injected cells (
Fig. 5F shows the mean current-voltage curves of chloride
currents obtained in antisense ClC-2 injected Caco-2 cells before and
after application of hypotonic shock. The average current density at
In a Polarized Caco-2 Cell Monolayer, ClC-2 Localizes Close to the
Tight Junction Complex--
To determine whether ClC-2 protein
exhibits a polarized distribution in fully differentiated intestinal
cells, we examined its subcellular localization in confluent Caco-2
cells grown on semipermeable filters. It has been well documented that
Caco-2 cells grown on either filters or on glass exhibit a polarized phenotype 4-6 days after plating (8, 9). In such a confluent monolayer, we found that ClC-2 protein exhibited a novel localization pattern.
Fig. 6 shows optical sections obtained
using a confocal microscopy of confluent Caco-2 cells co-labeled with
the polyclonal anti-ClC-2 antibody and a monoclonal antibody against
the transmembrane tight junction protein, occludin (22). The first row
of images shows the most apical sections, and the subsequent rows show
consecutive images obtained toward the basolateral pole. The ClC-2
signal (red) clearly delineates the plasma membrane in the
first row (Fig. 6), whereas the occludin signal (green) is
quite faint, suggesting that ClC-2 protein expression is apical
relative to this tight junction protein. The yellow signal, indicative
of regions of overlap, is weak in this apical section. In the next row
of images, the membrane staining corresponding to both proteins and the
yellow signal is intense, suggesting that expression of these proteins
may overlap in this optical section. The last two rows of sections show
the disappearance of the ClC-2 signal from the plasma membrane while
the occludin signal remains strong (Fig. 6). Thus, it appears that
ClC-2 localization overlaps with the apical aspect of the tight
junctions. Identical localization of ClC-2 was observed for confluent
monolayers of Caco-2 cells grown on glass coverslips (data not
shown).
ClC-2 Contributes to Hypotonicity-stimulated Chloride Secretion in
Monolayers of Caco-2 Cells--
There may be multiple functional
consequences of this unique localization of ClC-2 because the tight
junction is known to be critical for regulating solute (including ions)
and water flux through the paracellular pathway and as well maintaining
the polarization of membrane proteins and lipids (22, 23). We reasoned
that ClC-2 may contribute to chloride flux in a secretory direction across the apical membrane because its localization not only overlaps with the junction transmembrane protein, occludin, but it also extends
into the membrane at the apical aspect of the tight junction. Typically, chloride channels implicated in secretion have been localized to the apical membrane (24). Hence, we investigated the role
of ClC-2 in chloride secretion across monolayers of Caco-2 cells.
The application of hypotonicity to the mucosal or apical surface of
intestinal epithelia has been previously shown to stimulate chloride
secretion (25, 26). However, the molecular basis for this secretory
response remained unclear. To determine whether Caco-2 cell monolayers
model this response, we measured short circuit current responses to
hypotonic shock by Caco-2 cell monolayers mounted in Ussing chambers.
Hypotonicity-activated increases in luminally (or apically) directed
negative short circuit current (Isc) across
epithelia have been shown to correlate with chloride secretion in
previous studies Ussing chamber studies (25). Similarily, we found that
reducing the osmolarity of the bath facing the apical membrane of
confluent Caco-2 cells (by 20%) evoked a transient increase in
luminally directed, negative transepithelial short circuit current
(Isc) (Fig.
7A).
Isc increased significantly from
To determine directly whether ClC-2 contributes to this response, we
assessed the effect of antisense ClC-2 transfection. The above Ussing
chamber studies required that Caco-2 cell monolayers were grown on
semipermeable filter supports. We found that transfection of Caco-2
cells on such filters was not efficient compared with transfection of
monolayers on coverslips (see Western analysis on Fig. 4). Hence, we
utilized an iodide efflux method to monitor chloride secretion from
cells on glass coverslips. This method has been used extensively in the
study of chloride secretion through CFTR in epithelial monolayers (14,
15). In Fig. 7B, we show traces of the
time-dependent efflux of iodide from iodide loaded Caco-2
cell monolayers. Efflux from monolayers (each comprised of 1 × 106 cells) into control, iodide-free isotonic solutions
reflects efflux through constitutively open anion channels. The rate of iodide efflux is enhanced if monolayers are transferred to hypotonic (iodide-free) solutions (25% isotonicity) as shown in the traces in
Fig. 7B, presumably because of the activation of apically
localized chloride channels. The extent of iodide efflux measured
during the first minute has been plotted in the bar graph shown in Fig. 7B (right panel). We show that in mock
(vector-alone) transfected Caco-2 cell monolayers, the rate of iodide
efflux is much greater in monolayers exposed to hypotonic solutions
relative to the efflux rate from monolayers exposed to isotonic
solutions (p = 0.002). In antisense ClC-2 transfected
monolayers, the rate of iodide efflux in hypotonic solutions was
significantly reduced in comparison to the hypotonicity-evoked efflux
measured in monolayers transfected with vector alone (p = 0.001). This result suggests that ClC-2 contributes to
hypotonicity-activated anion secretion. On the other hand,
overexpression of ClC-2 does not affect the iodide efflux response in
hypotonic solutions (p = 0.33) but does appear to
increase the basal efflux (p = 0.009). Hence,
overexpression of ClC-2 confers an increase in basal anion secretion.
Hypotonic solutions do not cause a further increase in iodide efflux in cells overexpressing ClC-2, possibly because the high basal permeation rate quickly dissipates the driving force for further iodide flux.
The major aim of this study was to test the hypothesis that ClC-2
contributes to native chloride secretion by intestinal epithelia. In
support of this hypothesis, we provided evidence that ClC-2 is
endogenously expressed at the plasma membrane of Caco-2 cells where it
contributes to native currents. Immunofluorescence studies were used to
document cell surface localization and patch clamp studies established
that ClC-2-like currents can be detected. We used an antisense strategy
to confirm the identity of these native currents as ClC-2. Finally,
reduction of ClC-2 protein expression in Caco-2 cell monolayers by
ClC-2 antisense transfection resulted in a decrease in hypotonicity
evoked iodide efflux, a measure of chloride secretion, indicating that
ClC-2 can contribute to chloride secretion.
Although ClC-2-like currents have been previously described in
epithelial cells (27-29), the current work provides the first direct
molecular evidence to support a role for ClC-2 in chloride secretion by
differentiated epithelial cells. Caco-2 cells have been found to model
the functional properties of the human small intestine; hence, our
findings suggest that ClC-2 may contribute to chloride secretion by the
epithelium of this organ. Similarily, ClC-2 protein has been localized
to the apical membrane of rat neonatal respiratory epithelium (4, 5)
and may mediate secretion in this tissue. However, ClC-2 is not likely
contribute to chloride secretion in all transport epithelia.
Preliminary studies in our laboratory have localized ClC-2 to the
basolateral membrane of the mouse
colon2; hence, ClC-2 protein
is likely to mediate chloride reabsorption by this tissue.
The mechanisms for activation of ClC-2 channel function in
situ are currently unclear. In neurons, ClC-2 is basally active at
resting membrane potentials (2). The results of the present experiments
suggest that in epithelial cells, ClC-2 channels are partially active
at resting membrane potentials. Patch clamp studies of Caco-2 cells
revealed that chloride currents associated with ClC-2 expression,
i.e. those endogenous currents inhibited by ClC-2 antisense
and augmented by expression of exogenous ClC-2 expression, are
activated by membrane hyperpolarization to potentials more negative
than Several previous studies have suggested that luminal hypotonicity
induces transepithelial chloride secretion. A study on
Necturus enterocytes has shown that a swelling-activated
chloride conductance is present in the apical membrane of these cells
(32). In addition, cell swelling may stimulate transepithelial chloride
secretion in airway epithelium (33), the T84 colonic
epithelial cell line (34, 35), HT-29Cl.19A intestinal epithelial cells
(36), and rat ileum (25). The molecular identity of the chloride
channels contributing to this hypotonicity-activated secretory chloride conductance in these diverse epithelial tissues has not been
determined. Our current studies showing that antisense ClC-2
transfection reduced this function suggests that ClC-2 should be
considered as a candidate. However, we cannot rule out the possibility
that antisense ClC-2 transfection may have caused primary and/or
secondary changes in the expression of other proteins over the 48-h
transfection time period. It is conceivable that expression of
undefined paralogs of human ClC-2 may have been reduced directly by the
antisense rodent ClC-2 construct employed in the current studies.
Further, although the expression of another channel implicated that in hypotonicity-activated ion conductance ClC-3 (37, 38) did not change,
it remains possible that the expression of other unidentified hypotonicity-activated membrane proteins may have altered as a secondary response to antisense ClC-2 transfection.
Our confocal studies of ClC-2 protein localization in confluent Caco-2
cells indicated that upon cell differentiation, ClC-2 protein is
predominantly situated at the apical aspect of the tight junctions.
This distinctive localization for ClC-2 has also been detected in
murine intestinal tissue, and immunogold studies showed that ClC-2
resides primarily in the membrane at this site (12). This distribution
pattern is not typical for ion channels that have been implicated in
chloride secretion. For example CFTR, the chloride channel thought to
mediate chloride transport in many epithelial tissues, resides on the
brush border membrane (24). Future studies are required to determine
the molecular mechanisms that traffic ClC-2 to the apical aspect of the
tight junction and those mechanisms that may act to retain the channel at this site. Finally, we have yet to determine whether there is a
particular physiological significance for the concentration of an anion
channel at the apical aspect of the tight junction. It is well known
that the tight junction functions as a size and charge selective gate
restricting the paracellular transit of organic and inorganic solutes
(22, 23). Although our studies suggest that ClC-2, at the apical aspect
of this junction, contributes to chloride secretion, its unique
localization may also function to regulate the gate functions of the
tight junction.
An understanding of the molecular basis for chloride secretion by
epithelial tissues is key to identification of future therapies for
intestinal secretory diseases such as diarrheal diseases and cystic
fibrosis. Our studies indicate that ClC-2 contributes to the native
secretory capacity of intestinal tissue. Hence, modification of its
function or location may affect the severity of secretory diseases.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-minimum essential medium (Wisent Inc.) containing
10% fetal calf serum, with 2 mM glutamine, 100 units
penicillin G, and 100 µg/ml streptomycin sulfate at 37 °C in an
atmosphere of 5% CO2, 95% air. For patch clamp studies,
cells were used 1-2 days after plating onto 35-mm coverslips (Fisher).
For Ussing chamber studies of chloride currents and assessment of ClC-2
localization by confocal microscopy, Caco-2 cells were seeded at high
density (1 × 106 cells ml
1, 500 µl/filter) and grown to confluency on clear Snapwell (Costar) filters
(pore size, 4 µm; diameter, 12 mm). Filters were cultured at 37 °C
in an atmosphere of 5% CO2, 95% air, for 2 weeks with medium replacement every 2-3 days. The formation of an intact monolayer was assessed by measuring transepithelial resistance in
Ussing chambers. Transepithelial resistance was calculated using Ohm's
law, from measurements of the change in short circuit current measured
(Isc, µA) upon passing 1 mV across the
epithelium. Monolayers were considered acceptable when the
transepithelial resistance exceeded 500 Ohms/cm2 and the
transepithelial potential difference exceeded 2 mV. Alternatively, for
iodide efflux studies and assessment of protein expression by Western
blotting and confocal microscopy following transfection, Caco-2 cells
were grown for 4-6 days on glass coverslips to achieve a
differentiated phenotype, as documented by Sood et al.
(9).
70 °C with one intensifying screen.
-actin (Sigma, anti-
-actin clone AC-74) was used
at 1/1000 dilution. Immunoreactive protein was detected using the ECL
system (Amersham Pharmacia Biotech).
when filled with pipette solution (see below for composition).
Whole cell currents were measured using an Axopatch 200A patch clamp
amplifier (Axon Instruments, Foster City, CA) and were filtered at 100 Hz with a 6-pore Bessel Filter. Sampling rate was 4 kHz for most data,
and junction potentials were corrected. Voltage clamp protocols were
generated using pCLAMP software (version 7, Axon Instruments) via a
Pentium II computer interfaced with a 1200 series Digidata (Axon
Instruments) The same software package was used both for data
acquisition and analysis. Current-voltage relationships were determined
in a stepwise clamp protocol. From a holding potential of
30 mV,
voltage pulses of 3.0 s were applied from
160 to +40 mV in 20-mV
increments. The bath solution contained 140 mM
N-methyl-D-glutamine (NMDG) chloride, 2 mM MgCl2, 2 mM CaCl2, 5 mM HEPES, whereas the pipette solution contained 140 mM NMDG chloride, 2 mM MgCl2, 2 mM EGTA, and 5 mM HEPES. Both pipette and bath
solutions were adjusted to pH 7.4 and 260 mOsm. In experiments in which
the response to hypotonic shock was studied, the bath solution
contained 110 mM NMDG chloride, 2 mM
CaCl2, 2 mM MgCl2, 5 mM
HEPES, pH 7.4, and the osmolarity was adjusted to 303 mOsm with
sucrose. The hypotonic bath solution was made as above, maintaining
equal ionic strength and pH, except that the osmolarity was adjusted to
228 mOsm with sucrose as assessed using a 5500 Vapor pressure osmometer
(Wescor, Johns Scientific Inc.). Caco-2 cells were subjected to
hypotonic shock using a gravity-fed superfusion system.
) vector such that the reversed restriction sites on
this vector would reverse the orientation of the open reading frame to
create the antisense plasmid. The antisense murine ClC-4 construct
(Clcn4, a gift from E. Rugarli, Milano, Italy) was made by
cloning the ClC-4 open reading frame with BamHI (5') and
EcoRI (3') into pCDNA 3.1(
). The murine ClC-4 construct shares 73% sequence identity with the human sequence.
cm2. Filters were inserted into Ussing
chambers and bathed in a buffer composed of 110.4 mM NaCl,
27.5 mM mannitol, 2.4 mM
K2HPO4, 0.8 mM
KH2PO4, 10 mM glucose, 10 mM HEPES, and 1 mM CaCl2, gassed with 95% O2 and heated to 37 °C. After 5-8 min of
measuring basal current the apical solution was changed to one lacking
(27.5 mM) mannitol for 20% hypotonic shock (HTS; 80%
isotonicity; determined using a 5500 Vapor pressure osmometer, Wescor,
Johns Scientific Inc.).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
ClC-2 message and protein expression in
Caco-2 cells. A, Northern analysis shows that the ClC-2
cDNA probe recognizes a band of ~ 4.6 kb in Caco-2 cells.
The 6.2-kb marker is CFTR, and the 4.6- and 1.8-kb markers indicate the
position of ribosomal RNA. B, immunoblot analysis shows that
the polyclonal anti-ClC-2 antibody generated against a peptide within
the amino terminus of the rat ClC-2 sequence (residues 34-71) (11)
recognizes a broad band that corresponds to molecular mass of ~97 kDa
in Caco-2 cells (first lane). This 97-kDa band is competed
with 1.2-fold excess of the antigenic fusion peptide (ppt,
second lane) but not with GST alone (third lane),
confirming antibody specificity.
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Fig. 2.
Immunolocalization of ClC-2 in the plasma
membrane of Caco-2 cells. A, image of ClC-2
localization in Caco-2 monolayer detected by immunofluorescence using
an epifluorescence microscope (see "Experimental Procedures").
B, the immunofluorescence specifically labels ClC-2 as it
can be competed by preincubation with 1.9-fold excess of the ClC-2
fusion protein (ppt, right panel). Images were
obtained using a 40× objective.
30 mV, the membrane potential was stepped by
20-mV increments from
160 to + 40 mV. Because ClC-2 currents have not
been reported to be ATP-dependent and to minimize the
contribution by the ATP-dependent, swelling-activated outwardly rectifying chloride channel, volume-sensitive organic osmolyte anion channel (17), MgATP was not included the patch pipette
solutions. As shown in Fig. 3
(A and C), currents typical of those previously
associated with ClC-2 expression, i.e. showing activation
with hyperpolarizing voltage steps and an inwardly rectifying
current-voltage relationship, were detected in Caco-2 cells. At the
hyperpolarized membrane potential of
160 mV, these currents had a
magnitude of
37 pA/pF ± 0.96 (n = 8). These
hyperpolarization-activated currents reversed close to the estimated
equilibrium potential of chloride (ECl = 0 in
symmetrical NMDG chloride solutions (Fig. 3C,
circles).
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Fig. 3.
ClC-2-mediated chloride currents are
activated by hypotonic shock in Caco-2 cells. ClC-2 mediated
chloride currents recorded in Caco-2 cells before (A) and
after (B) exposure to 25% hypotonic shock. The same voltage
step protocol was applied as described previously. C, mean
I/V curves of ClC-2 current under isotonic
(circle, n = 4) and hypotonic (square,
n = 4) conditions.
38 pA/pF to
84 pA/pF
within 3-5 min after bath dilution at
160 mV (Fig. 3). The mean
current-voltage (I/V) curves of ClC-2 currents
before and after application of hypotonic shock reversed close to the
chloride equilibrium potential (+5.6 ± 0.47 mV) as shown in Fig.
3C. The average current at
160 mV following hypotonic
shock (
65 ± 8 pA/pF) was elevated when compared with currents
measured in isotonic condition (
32 ± 2 pA/pF)
(p = 0.0208). The I/V
relationship of the HTS-stimulated chloride currents was less inwardly
rectifying than that observed in isotonic solutions (Fig.
3C). A similar change in the I/V
relationship was observed with HTS in chloride currents specifically
conferred by ClC-2 expression in Xenopus oocytes and
Sf9 cells and has been attributed to an alteration in the
inactivation gate of ClC-2 (11, 16). These results indicate that native
ClC-2 expression at the cell surface of Caco-2 cells is associated with
appearance of chloride currents with activation and conductance
properties similar to those conferred by ClC-2 expression in
heterologous expression systems (1, 11, 16, 18).
-actin expression that differences in protein loading could not account for the decrease in ClC-2 expression in the antisense transfected cells (Fig. 4A).
Furthermore, we examined the effects of antisense ClC-2 transfection on
immunolabeled ClC-2 detected by fluorescence confocal microscopy. DNA
coding for green fluorescence protein (GFP) was cotransfected with
antisense ClC-2 (or empty vector as a control) into Caco-2 cells to
identify transfected cells (Fig. 4B). We used an imaging
program (Scion Corp.) to compare the ClC-2 immunofluorescence intensity
in antisense ClC-2 and in vector transfected Caco-2 cells. We found
that the fluorescence intensity of the signal (red)
corresponding to membrane expression of ClC-2 was reduced by ~75% in
antisense ClC-2 transfected Caco-2 cells (24.9 units ± 4.3, n = 13, p < 0.0001) relative to the
intensity of the ClC-2 signal in mock transfected cells (105.8 units ± 3, n = 10). Immunofluorescence
corresponding to expression of ClC-3, a related family member, was not
affected by antisense ClC-2 transfection (Fig. 4C). The
signal detected using this ClC-3 antibody in immunofluorescence studies
can be competed using the antigenic peptide used to raise the antibody,
confirming its specificity (19). Fig. 4C shows that the
ClC-3 immunofluorescence (red) in antisense ClC-2 and GFP
cotransfected Caco-2 cells (106.1 units ± 3.2, n = 20) was similar to that in vector and GFP cotransfected cells (105.9 units ± 2.2, n = 17, p = 0.97).
Interestingly, our studies show that unlike ClC-2, immunoreactive ClC-3
appears to be primarily expressed in intracellular membranes, although
there is signal detected at the cell surface in a subpopulation of
cells.
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Fig. 4.
ClC-2 antisense reduces ClC-2 protein
expression in Caco-2 cells. A, Western analyses show
reduced ClC-2 expression in ClC-2 antisense transfected monolayers of
Caco-2 cells. 50 µg of protein were loaded per lane. -actin
labeling confirms that the reduction of ClC-2 signal does not reflect
less sample. B, upper panels show confocal image
of immunolabeled ClC-2 (red) endogenously expressed in
Caco-2 cell cotransfected with cDNA coding for empty vector
(Vtr) and GFP. Lower panels, the
immunofluorescence corresponding to ClC-2 signal in antisense ClC-2 and
GFP cotransfected Caco-2 cells. Images were obtained using a 63×
objective. Fluorescence intensity of the ClC-2 signal was quantitated
by averaging the pixel intensity of the grayscale image (0 units = white, 255 units = black) of eight regions
in the plasma membrane. These points on the membrane were assigned
using four lines which bi-sect each other, and each line intercepts the
membrane two times. The bar graph shows the means ± S.E. of fluorescence intensities corresponding to ClC-2 membrane
expression for vector (n = 10) and antisense ClC-2
transfected (aClC-2, n = 13) Caco-2 cells.
C, the upper panels show confocal image of
immunolabeled ClC-3 (red) endogenously expressed in Caco-2
cell cotransfected with cDNA coding for empty vector and GFP.
Lower panels, the immunofluorescence corresponding to ClC-3
signal in antisense ClC-2 and GFP cotransfected Caco-2 cells.
Fluorescence intensity of the ClC-3 signal was quantitated by averaging
pixel intensity of the grayscale image of eight randomly selected
regions around the nucleus delimited using four lines transecting the
nucleus. The bar graph shows the means ± S.E. of
fluorescence intensity determined in vector (n = 17)
and antisense ClC-2 transfected (n = 20) Caco-2
cells.
160 mV decreased from
37
pA/pF ± 1 (n = 8) in uninjected cells to
26 ± 2 pA/pF (n = 5, p = 0.0003) and
12 ± 1 pA/pF (n = 10, p < 0.0001) in 50 and 300 µg/ml antisense ClC-2
cDNA injected cells, respectively (Fig. 5, A and
B). To allow direct comparison of current amplitude in ClC-2
antisense injected and uninjected Caco-2 cells, we normalized these
currents to the currents at
160 mV in uninjected cells (Fig.
5B). As shown in Fig. 5B, the normalized ClC-2
currents in cells injected with 50 or 300 µg/ml antisense plasmid
were decreased by 29 ± 6 and 68 ± 2%, respectively.
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Fig. 5.
ClC-2 antisense reduces hyperpolarization and
hypotonic activated chloride currents in Caco-2 cells.
A, hyperpolarization activated chloride currents are reduced
in Caco-2 cells microinjected (intranuclear) with antisense ClC-2
plasmid (300 µg/ml). Whole cell currents obtained by voltage steps of
20 mV increments, applied from 160 mV to +40 mV. Initial holding
potential,
30 mV; final holding potential,
60 mV. The patch pipette
and bath solutions both contained NMGD chloride solutions (see
"Experimental Procedures" for more detail on buffers).
B, mean current-voltage relationship for chloride currents
obtained from control Caco-2 cells (n = 8, Ctl) from cells injected with 50 µg/ml ClC-2 antisense
(n = 5, aCLC-4) or 300 µg/ml ClC-2
antisense (n = 10). Currents were normalized to cell
capacitance (pF). The bar graph shows mean
currents in control and ClC-2 antisense microinjected Caco-2 cells at
160 mV. Currents were normalized to the mean current measured in
uninjected cells at
160 mV. C, mean
I/V curves of hyperpolarization-activated
chloride currents in antisense ClC-4 injected (square,
n = 4) and in noninjected (circle, n = 4) Caco-2 cells. ClC-2 currents were comparable in both uninjected and
ClC-4 antisense injected Caco-2 cells (p = 0.1849).
D, mean I/V curves in Caco-2 cells
microinjected with ClC-2 cDNA in the sense orientation
(square, n = 4) and in noninjected
(circle, n = 4) Caco-2 cells.
Hyperpolarization-activated currents (at
160 mV) are elevated in
ClC-2 cDNA microinjected cells when compared with control
(p = 0.0002). E, mean
I/V curves of ClC-2 current under hypotonic
condition in antisense ClC-2 injected (square,
n = 7) and in noninjected (circle,
n = 4) Caco-2 cells. Antisense ClC-2 expression
significantly diminished chloride currents measured at
160 mV and +40
mV. *, p < 0.0001; **, p = 0.0053. F, mean I/V curves of ClC-2 current in
antisense ClC-2-injected Caco-2 cells before (Iso,
square, n = 7) and after (Hypo,
circle, n = 7) exposure to 25% hypotonic
shock are shown. Currents measured at
160 mV in isotonic conditions
are not significantly different than currents measured in hypotonic
solutions (p = 0.26), but a significant difference with
hypotonic solutions was detected at +40 mV (p = 0.006).
31 ± 3 pA/pF at
160 mV, n = 4)
was not significantly different from that measured in noninjected cells
(
35 ± 2 pA/pF, p = 0.1849). Fig. 5D
shows that the amplitude of the hyperpolarization-activated, inwardly
rectifying chloride current was doubled by expression of exogenous
ClC-2 cDNA (
66 ± 3 pA/pF, n = 4). Together,
these results indicate that ClC-2 natively expressed in Caco-2 cells mediates inwardly rectifying, hyperpolarization-activated chloride currents.
15.7 ± 2 pA/pF, n = 7, p < 0.0001). The current density at +40 mV after HTS
was 12.7 ± 2.4 pA/pF in control cells and almost two to three
times less in antisense-transfected cells (4.9 ± 0.8 pA/pF,
p = 0.0053), suggesting that ClC-2 mediates most of the
hypotonicity-activated current at both potentials.
160 mV following hypotonic shock (
15.7 ± 2 pA/pF, n = 7) was not significantly different when compared
with current densities measured in isotonic conditions (
13.7 ± 0.9 pA/pF, n = 7, p = 0.26). Hence,
antisense ClC-2 transfection abolished hypotonicity activated chloride
currents at
160 mV. At +40 mV, current density determined in
antisense-transfected cells was less than half that measured in control
cells, confirming that ClC-2 is a major determinant of
hypotonicity-activated chloride currents evoked in these particular
experimental conditions, i.e. without MgATP added to the
pipette solutions. However, antisense ClC-2 did not abolish the
hypotonicity-evoked chloride currents at +40 mV, suggesting that
channels other than ClC-2 may also contribute to this response at
depolarized membrane potentials.
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Fig. 6.
Localization of ClC-2 protein to the apical
aspect of the tight junctions formed between Caco-2 cells in a
confluent monolayer. Consecutive optical cross-sections of
confluent Caco-2 cells that have been co-labeled with the polyclonal
anti-ClC-2 antibody (red) and a monoclonal anti-occludin
antibody (green) were obtained by confocal microscopy.
Panels 1-5 extends from the apical toward the basolateral
membrane. Panel 2 is the most apical section in which the
ClC-2 signal could be detected, and panel 5 is the last
section in which membrane staining of ClC-2 could be detected. Pictures
were taken using a confocal microscope with a 100× objective.
1.0 ± 0.1 µA/cm2 to
3.1 ± 0.4 µA/cm2
(n = 14, p < 0.0001, Student's test
for paired data) with this treatment. These findings are consistent
with the transient activation of an apical chloride conductance path by
luminal hypotonicity in Caco-2 cells.
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Fig. 7.
Application of hypotonic shock to apical
membranes of confluent Caco-2 cell monolayers stimulates chloride
secretion through ClC-2 channels. A, left
panel, Caco-2 cell monolayers grown on semipermeable support were
mounted in Ussing chamber to assess short circuit current response to
dilution of mucosal solution. A transient increase in negative
Isc was activated by exchange of the isotonic
mucosal bath with 20% hypotonic mucosal bath as indicated by the
bar. Right panel, the mean negative basal
transepithelial Isc was increased by hypotonic
shock (open bars, n = 14) (p < 0.05, Student's paired test). B, left panel,
iodide efflux from Caco-2 monolayers transfected with pCDNA 3.1(+)
vector alone (Vtr) or antisense ClC-2 construct was
monitored continuously using an iodide sensing electrode. Monolayers
were exposed either to isotonic (Ctl) or 25% hypotonic
solutions. Right panel, this bar graph shows the
means ± S.D. iodide efflux measurements (iodide efflux during the
first minute) from vector-alone transfected and antisense ClC-2
transfected Caco-2 cell monolayers as well as monolayers transfected
with ClC-2 in the sense orientation. The solid bars
represent data from isotonic solutions, and the open bars
represent data from hypotonic solutions. A minimum of four monolayers
were studied for each condition.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
60 mV. This membrane potential is close to the resting membrane
potential cited for gastrointestinal epithelial cells, i.e.
from
50 to
60 mV (26). Gastrointestinal epithelial cells can,
however, reach more hyperpolarized potentials when stimulated by the
hormones that act to increase potassium permeability, i.e.
acetylcholine and vasoactive intestinal peptide (26, 30). The
acetylcholine analogue, carbachol, has been reported to cause transient
hyperpolarizations of 10-25 mV in isolated small intestinal crypts.
Hence, during stimulation with the above hormones, ClC-2 channels may
become further activated. In the present studies we diluted the
external solutions to 70-80% isotonicity to observe stimulation of
ClC-2-mediated currents in single cells and iodide efflux from Caco-2
cell monolayers. It remains to be determined whether this experimental
maneuver reflects a physiologically relevant stimulus to intestinal
epithelial cells such as the generation of osmotic gradients during the
concentrative uptake of nutrients (31).
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ACKNOWLEDGEMENTS |
---|
We are grateful to Dr. T. Jentsch for the gift of rat ClC-2 cDNA and to Dr. E. Rugarli for the gift of mouse ClC-4 cDNA. We also acknowledge the helpful discussions with Dr. Herman Yeger at the Hospital for Sick Children, Toronto.
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FOOTNOTES |
---|
* This work was funded by a National Institutes of Health grant (to C. E. B.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
These authors contributed equally to this work.
§ Recipient of a Fellowship award from the Canadian Cystic Fibrosis Foundation.
¶ Recipient of a Studentship award from the Canadian Cystic Fibrosis Foundation.
To whom correspondence should be addressed: Research Inst.,
Hospital for Sick Children, 555 University Ave., Toronto, Ontario M5G
1X8, Canada. Tel.: 416-813-5981; Fax: 416-813-5028; E-mail: bear@ sickkids.on.ca.
Published, JBC Papers in Press, November 28, 2000, DOI 10.1074/jbc.M006764200
2 K. Gyömörey and C. Bear, unpublished data.
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ABBREVIATIONS |
---|
The abbreviations used are: CF, cystic fibrosis; CFTR, CF transmembrane conductance regulator; GST, glutathione S-transferase; NMDG, N-methyl-D-glutamine; HTS, hypotonic shock; kb, kilobase(s); GFP, green fluorescence protein.
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REFERENCES |
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