1 Cystic Fibrosis Research Center and 2 Center for Biologic Imaging, Department of Cell Biology and Physiology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania 15261
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ABSTRACT |
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Mutations in the gene encoding the cystic fibrosis (CF) transmembrane conductance regulator (CFTR) chloride channel give rise to the most common lethal genetic disease of Caucasian populations, CF. Although the function of CFTR is primarily related to the regulation of apical membrane chloride permeability, biochemical, immunocytochemical, and functional studies indicate that CFTR is also present in endosomal and trans Golgi compartments. The molecular pathways by which CFTR is internalized into intracellular compartments are not fully understood. To define the pathways for CFTR internalization, we investigated the association of CFTR with two specialized domains of the plasma membrane, clathrin-coated pits and caveolae. Internalization of CFTR was monitored after cell surface biotinylation and quantitation of cell surface CFTR levels after elution of cell lysates from a monomeric avidin column. Cell surface levels of CFTR were determined after disruption of caveolae or clathrin-coated vesicle formation. Biochemical assays revealed that disrupting the formation of clathrin-coated vesicles inhibited the internalization of CFTR from the plasma membrane, resulting in a threefold increase in the steady-state levels of cell surface CFTR. In contrast, the levels of cell surface CFTR after disruption of caveolae were not different from those in control cells. In addition, although our studies show the presence of caveolin at the apical membrane domain of human airway epithelial cells, we were unable to detect CFTR in purified caveolae. These results suggest that CFTR is constitutively internalized from the apical plasma membrane via clathrin-coated pits and that CFTR is excluded from caveolae.
Calu-3 cells; caveolin; endocytosis; ion channel
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INTRODUCTION |
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CYSTIC FIBROSIS (CF) is the most frequently occurring recessive genetic disorder of Caucasian populations, affecting 1 in 2,500 live births. The CF gene encodes an integral membrane glycoprotein, the CF transmembrane conductance regulator (CFTR) (34), which functions as a chloride channel within the apical membrane of polarized epithelial cells (1, 2, 19). Alterations in the gene encoding CFTR give rise to a characteristic phenotype in CF cells, namely, impaired transepithelial chloride secretion in response to activation of the cAMP-mediated second messenger cascade (23, 38). Data from patch-clamp studies (1, 19) suggest that plasma membrane-associated CFTR can be directly regulated via a phosphorylation-dephosphorylation mechanism. However, an implicit prerequisite for this simple model is the continued presence of an appropriate number of CFTR channels in the apical membrane to sustain the required level of chloride secretion. Studies by Bradbury and colleagues (5, 6) focused on the hypothesis that regulation of the number of active chloride channels in the apical membranes of epithelial cells can, in part, be brought about by the regulated insertion and internalization of CFTR into and from the plasma membrane. Indeed, both immunologic (17, 33) and electrophysiological (44) studies have reported increases in the copy number of plasma membrane CFTR and apical membrane surface area in response to forskolin. Such alteration in copy number in plasma membrane transporters in response to stimulation has been reported for a growing number of transporters including the insulin-responsive glucose transporter GLUT-4 and the antidiuretic hormone-responsive water channel (for a review see Ref. 3).
Important to an understanding of the molecular mechanisms involved in
CFTR translocation is knowledge of the proteins that localize with CFTR
in the same compartment. Recent studies have identified some of the
components involved in the vesicle trafficking of CFTR, including
-adaptin and clathrin heavy chain (4), rab4 (47), and syntaxin 1A (28, 31).
The coexistence of several different internalization pathways has been
documented in many cell types (22), including clathrin-dependent and
-independent endocytosis, macropinocytosis, and potocytosis. Bradbury
et al. (4) previously detected CFTR within clathrin-coated vesicles; however, whether this is the sole route for CFTR internalization from
the plasma membrane remains to be determined.
Caveolae are small, uniform, -form invaginations of the cell
surface, having a diameter of 50-80 nm. Caveolae have been
identified in many different cell types including smooth muscle cells,
adipocytes, fibroblasts, and epithelial cells (9, 14). Several workers have investigated the relevance of caveolae to the regulated
trafficking of integral membrane transport proteins, including GLUT-4
and the vacuolar proton ATPase of renal intercalated cells (8). For
example, although some workers (37) have suggested that caveolae may
play a part in the trafficking of GLUT-4, others (18, 27) have been
unable to document a relationship between caveolae and the molecular
machinery involved in the trafficking of GLUT-4. The aim of our present
study was to evaluate the distribution and expression of caveolin (a
generally accepted marker for caveolae) in a variety of chloride
secretory epithelial cell lines and to determine the roles of caveolae
and clathrin-coated vesicles as pathways for the internalization of
CFTR from the plasma membrane.
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METHODS |
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Materials. Antibodies to
caveolin-1, -2, and -3, clathrin heavy chain, and protein kinase (PK)
C- were obtained from Transduction Laboratories (Lexington, KY).
Monoclonal antibodies to CFTR were from Genzyme (Framingham, MA).
Antibodies to ZO-1 were from Zymed Laboratories (San Francisco, CA).
Optiprep and cell culture media were from GIBCO BRL (Grand Island, NY).
All other materials were obtained from Sigma (St. Louis, MO) and were
of reagent grade quality. All solutions were prepared in Milli-Q water
and filtered (0.45 µm) before use.
Cell culture. C127-CFTR, CFPAC-PLJ-CFTR, and HeLa cells were grown in DMEM containing 10% fetal bovine serum (FBS). Calu-3 cells were grown on filters at an air interface in DMEM-Ham's F-12 medium containing 15% FBS. T84 cells were grown in DMEM-Ham's F-12 medium containing 5% heat-inactivated FBS. HT-29 and Caco-2 cells were grown in DMEM containing 10% FBS. All cell lines were maintained on 100-mm plastic culture dishes. Cells were grown to confluence before use, usually 7-10 days postplating.
Electrophoresis and immunoblot. Cells
solubilized in Laemmli sample buffer containing 1.2%
-mercaptoethanol were heated to 37°C for 10 min before being
loaded onto gels. The proteins were resolved by SDS-PAGE and
transferred to nitrocellulose. The nitrocellulose was blocked (5%
nonfat dry milk in 10 mM Tris, pH 7.5, 100 mM NaCl, and 0.1% Tween 20)
for 1 h at room temperature and incubated overnight at 4°C with
primary antibodies. The nitrocellulose was washed extensively and
incubated with secondary antibodies (rabbit anti-mouse or goat
anti-rabbit conjugated to horseradish peroxidase) for 1 h at room
temperature. The nitrocellulose was washed again, and the bands were
visualized with enhanced chemiluminescence.
Fluorescence microscopy of caveolin-1 expression. To evaluate the expression of caveolin-1 within cells, standard immunohistochemical methods were used. Cells were lightly fixed in 2% paraformaldehyde in phosphate-buffered saline (PBS) for 10 min followed by a permeabilization step with 2% paraformaldehyde containing 0.1% Triton X-100 in PBS for 10 min. The cells were then washed three times in PBS containing 0.5% BSA and 0.15% glycine, pH 7.4 (buffer A) followed by a 30-min incubation with purified goat IgG (50 µg/ml) at 25°C and three additional washes with buffer A. All the preceding steps were designed to ensure minimal nonspecific reaction to the antibodies used. The cells were then incubated for 4 h with a mixture of two primary antibodies (a monoclonal IgG1 directed against caveolin-1 and a polyclonal antibody directed against ZO-1) followed by three washes in buffer A and a 2-h incubation in a mix of fluorescently labeled secondary antibodies (an indocarbocyanine-conjugated anti-mouse IgG to detect the caveolin-1 antibody and an FITC-conjugated anti-rabbit IgG to detect the ZO-1 antibody). The cells were then washed six times (5 min/wash) in buffer A and then mounted in Gelvatol, and a coverslip was applied for light microscopy. Observation was with a Zeiss Axiovert 135 microscope equipped with epifluorescence optics, computer-controlled shutters, stage control, and cameras. ZO-1 was visualized with an FITC filter combination (Chroma) and caveolin-1 with a indocarbocyanine filter set (Chroma). Images were collected with a cooled charge-coupled device camera (Photometrics) at 1,300 × 1,000 pixel resolution with a ×100 1.3-numerical aperture plan apochromatic lens. Microscope control and image collection were managed by ONCORimage (ONCOR, Gaithersburg, MD). To generate deconvolved profiles of optical sections throughout the cells, serial image planes were collected throughout the depth of the cells at 0.2-µm separation. After image collection, the individual image stacks for caveolin-1 and ZO-1 were examined, and appropriate 512 × 512 subregions within the individual image profiles were selected. This was to ensure a reasonable time frame for deconvolution. Deconvolution was performed with an exhaustive photon reassignment algorithm on a Silicon Graphics Indigo2 workstation with XCOSM (University of Washington, Seattle). To generate rendered images, the resultant deconvolved stacks were ported into ImageSpace 3.2 (Molecular Dynamics), and image projections were generated with a look-through projection algorithm. Images were assembled in Photoshop 4.0 with no further processing or enhancement.
Detergent solubilization. The method of Brown and Rose (10) was used to prepare low-density, detergent-insoluble complexes. Cell monolayers were lysed with 1 ml of extraction buffer (25 mM HEPES, pH 7.5, 150 mM NaCl, and 1% Triton X-100) for 20 min on ice. Detergent-soluble and -insoluble fractions were collected by centrifugation. Detergent-soluble material was brought to 1× radioimmunoprecipitation assay (RIPA) buffer (50 mM Tris · HCl, pH 7.5, 150 mM NaCl, 1% Triton X-100, 1% sodium deoxycholate, and 0.1% SDS) by addition from a 5× stock. Detergent-insoluble material was resuspended in 800 µl of extraction buffer followed by the addition of 200 µl of 5× RIPA buffer. Detergent-soluble and -insoluble materials were then analyzed for caveolin-1 by immunoblot and for CFTR by immunoprecipitation.
Immunoprecipitation. CFTR was
immunoprecipitated as previously described (16) with a monoclonal
antibody to the carboxy terminus of CFTR. The immunoprecipitated CFTR
was phosphorylated with a cAMP-dependent PK catalytic subunit and
[-32P]ATP followed
by resolution on SDS-PAGE gels. After fixation, the gels were dried and
processed for autoradiography or phosphorimaging (Bio-Rad).
Biotinylation. Cell surface biotinylation was performed as previously described (32). Briefly, monolayers were washed in PBS at 4°C, then incubated with NaIO4 in the dark for 30 min at 4°C. The cells were again washed in PBS and labeled with biotin-LC-hydrazide (Pierce Chemical, Rockford IL). After being labeled, the cells were extensively washed in PBS and lysed in buffer (20 mM HEPES, pH 7.4, 150 mM NaCl, 1 mM EDTA, and 1% Nonidet P-40). Separation of biotinylated and nonbiotinylated proteins was performed with a column of immobilized monomeric avidin (Pierce Chemical) by applying equal amounts of cell protein to each column according to the manufacturer's instructions. Biotinylated proteins eluted in the presence of excess free avidin were subjected to immunoprecipitation with antibodies against CFTR as described in Immunoprecipitation. SDS-PAGE gels containing biotinylated CFTR were subjected to phosphorimage analysis with a Bio-Rad phosphorimager.
Determination of 125I-labeled transferrin uptake. A minor modification of the procedure described by Simon et al. (39) was used to analyze the binding and uptake of 125I-transferrin. 125I-transferrin was synthesized according to a previously published protocol by Simon et al. (39) to a specific activity of 0.5 µCi/mg. Cells grown to confluence at an air interface on Transwell filters were incubated with medium containing 1% BSA instead of 10% FBS for 1 h (to internalize any transferrin bound to surface receptors) and then cooled to 2°C. The cells were then incubated for 30 min in the same medium containing 10 µg/ml of 125I-transferrin, also at 2°C. To measure uptake, the cells were warmed rapidly to 37°C and then cooled to 2°C after 10 min. The cells were then washed at 2°C twice with medium-1% BSA, three times with PBS-1% BSA, and then four times with PBS. The cells were released from the filters by incubation for ~30 min in PBS containing 0.5% trypsin at 2°C, transferred to a microcentrifuge tube, and rotated in the same medium for ~45 min to ensure complete release of 125I from the cell surface. Serum (10%) was added to quench the trypsin, and the cells were centrifuged (10,000 g for 5 min) and washed by resuspension in PBS. The supernatant contained surface-bound 125I-transferrin, and the cell pellet, which contained internalized 125I-transferrin, was analyzed for radioactivity. When the effect of hypertonic exposure or K+ depletion was examined, transferrin uptake was measured in hypertonic DMEM-Ham's F-12 medium or K+-free medium.
Disruption of caveolae. Two different approaches were employed to disrupt caveolae in Calu-3 cells. 1) Cells were incubated in filipin (1 µg/ml) for 30 min at 37°C before isolation of a caveolae fraction in the continued presence of filipin or for CFTR immunoprecipitation. 2) Caveolae were also disrupted by incubating Calu-3 cells with cholesterol oxidase (0.5 U/ml) for 60 min at 37°C before CFTR immunoprecipitation. Inhibition of clathrin-dependent endocytosis. Three independent approaches were employed to inhibit clathrin-coated vesicle formation in Calu-3 cells. 1) Cells were incubated in chlorpromazine (100 µM) for 30 min at 37°C before CFTR immunoprecipitation. 2) Depletion of cytosolic K+ was achieved by hypertonic swelling of the cells in a 50% diluted DMEM-Ham's F-12 medium containing ouabain (1 mM) for 5 min and then incubating them in ice-cold K+-free medium (100 mM NaCl, 50 mM HEPES, 1 mM MgCl2, and 100 mM glucose, pH 7.3) for 20 min. 3) Cells were incubated in a hypertonic medium (DMEM-Ham's F-12 medium supplemented with 0.3 M sucrose) for 15 min at 37°C. Caveolae isolation. Caveolae were isolated with the method of Smart et al. (42). Briefly, cells were homogenized (20 mM Tricene, pH 7.8, 250 mM sucrose, and 1 mM EDTA) and centrifuged to yield a postnuclear supernatant. Plasma membranes were obtained by layering the postnuclear supernatant on top of 30% Percoll followed by centrifugation (84,000 g for 30 min). The plasma membranes were then disrupted by sonication, and the caveolae were isolated from other plasma membrane domains by flotation in a 20-10% linear Optiprep gradient. Protein determination. Protein concentrations were determined according to the method of Bradford (7), with BSA fraction V as a standard. ![]() |
RESULTS |
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Caveolin and CFTR show no correlation in tissue
expression. To investigate potential localization of
CFTR in caveolae, experiments were performed to determine whether CFTR
and caveolin, a generally accepted marker for caveolae, are expressed
in the same cell types. Immunoblot analyses were performed on a panel
of cultured epithelial cell lines endogenously expressing CFTR (Calu-3,
T84, HT-29, and Caco-2), cell lines stably expressing CFTR (C127-CFTR
and CFPAC-PLJ-CFTR), and cells not expressing CFTR (HeLa). Cell lysates
(25 µg) were resolved by SDS-PAGE, transferred to nitrocellulose, and
probed with various antibodies. Because Bradbury et al. (4) previously showed sequestration of CFTR into clathrin-coated
vesicles, we probed the cell lines for clathrin heavy chain (Fig.
1A,
Table 1). As expected,
clathrin heavy chain appeared as a single band of 180 kDa in all cell
lines tested, consistent with the ubiquitous role of clathrin in
membrane and solute internalization. Moreover, similar levels of
clathrin heavy chain were observed in all cell lines tested. In
contrast, immunoblot analyses for caveolin revealed a different pattern
of expression. Caveolin-1 in the mouse epithelioid cell line C127-CFTR
appeared as a 20- to 22-kDa doublet as reported for
several other cell lines (12, 21, 25). Caveolin-1 was also detected as
a single band in a human pancreatic ductal cell line (CFPAC-PLJ-CFTR),
a model of human airway serous epithelial cells (Calu-3), and
a human colonic epithelial cell line (HT-29). A human endothelial cell
line (HeLa) and models of human colonic epithelia (T84 and Caco-2) were
devoid of caveolin-1 (Fig. 1A, Table
1). Even for those cells expressing caveolin-1, the level of expression
varied considerably between cell lines. Caveolin-2 appeared as a
band of 20 kDa in C127-CFTR, HeLa, and CFPAC-PLJ-CFTR cells and at low levels in Calu-3 cells (Fig.
1A, Table 1). Caveolin-2 was not
present in T84, HT-29, or Caco-2 cells. As with caveolin-1, caveolin-2
expression levels varied between the cell lines tested. Caveolin-3, a
muscle-specific caveolin isoform, was not detected in any of the cell
lines tested (caveolin-3 was detected in a rat muscle cell lysate; data
not shown). Several signaling molecules have been reported to be
associated with or enriched in caveolae, including PKC- (42).
Immunoblot analyses on cell lysates revealed the presence of
PKC-
in all cell lines tested as a single band of 82 kDa (Fig.
1A, Table 1).
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The above panel of cell lines was also subjected to immunoprecipitation with an antibody directed against CFTR. Mature fully glycosylated band C CFTR was observed as a band of ~170-180 kDa in C127-CFTR, Calu-3, T84, HT-29, and Caco-2 cells. CFPAC-PLJ-CFTR cells expressed low, but detectable, levels of CFTR, whereas HeLa cells were devoid of CFTR (Fig. 1B, Table 1). The level of CFTR expression varied between the cell lines tested, with the highest level of endogenous CFTR expression being seen in Calu-3 human airway serous cells. In addition, slightly different molecular masses for CFTR were observed, indicative of variations in glycosylation patterns between the various tested cell lines. The core-glycosylated band B CFTR occurred at the same molecular mass in all samples. As can be seen, the level of CFTR expression in the C127-CFTR-overexpressing system was much greater than that observed in natively expressing cells, being approximately six- to sevenfold higher than that observed in Calu-3 cells. No correlation between the expression of CFTR and the expression of caveolin-1, -2, or -3 was observed.
Caveolae are present in the apical domain of polarized
Calu-3 cells. For there to be any physiologically
relevant interaction between CFTR and caveolae, caveolae must be
present at the apical surface of polarized secretory epithelia (because
CFTR is an apical membrane ion channel). Because C127-CFTR and Calu-3
cells express high levels of both caveolin and CFTR, we focused our
further studies on these cell lines. We verified the presence of
caveolin in the apical domain of polarized Calu-3 cells by
immunofluorescence microscopy followed by image analysis on a Silicon
Graphics workstation (Fig. 2). ZO-1 was
used as marker for tight junctions (Fig. 2, green signal) and
delineated the apical and basolateral domains of the cell. Optical
sections at and above the level of the tight junctions (Fig.
2B) show discrete punctate staining
for caveolin evenly distributed across the cell surface. Below the
level of the tight junctions (Fig.
3B),
very little caveolin-1 is observed. Figure
2A shows a composite reconstruction of
all optical sections through the Calu-3 cells, and Fig. 2,
B and
C, shows representative single optical
sections taken from this data set. Although most cells contain similar
levels of caveolin, a few cells express high levels of caveolin. The
observation that caveolin, and hence caveolae, are indeed present in
the apical membrane domain of polarized Calu-3 cells is consistent with
the finding that caveolin is present in both the apical and basolateral
domains of polarized MDCK cells (13, 25, 36) and allows for potential
interaction between CFTR and caveolae.
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Disruption of clathrin-coated vesicles, but not of
caveolae, alters the cell surface expression of CFTR.
To investigate whether CFTR internalization in polarized Calu-3
epithelial cells is mediated by clathrin-coated vesicles, caveolae, or
both, we monitored levels of biotinylated cell surface CFTR. This
approach has previously been utilized to monitor changes in cell
surface CFTR on activation of the cAMP-mediated second messenger
cascade (32). Inhibiting the internalization of CFTR from the cell
surface would be expected to lead to an accumulation of CFTR at the
plasma membrane, where it would be available for biotinylation. We
employed two independent methods to disrupt caveolae. First, we
utilized the sterol binding drug filipin (15, 29). As shown in Fig.
3A, exposure of Calu-3 cells to
filipin [1 µg/ml, a concentration previously shown to result in
maximum inhibition of fluid-phase endocytosis in endothelial cells
(15)] resulted in the disruption of caveolae as monitored by a
loss of caveolin-1 from relevant fractions of the Optiprep gradient.
Densitometric analysis revealed that exposure of Calu-3 cells to
filipin resulted in the loss of 92 ± 5% of the caveolin signal
from Optiprep-fractionated plasma membranes. Although caveolae were
disrupted, such disruption did not affect the internalization of cell
surface CFTR (Fig. 3C, Table
2). Thus the level of cell surface CFTR was not significantly different from that of control cells
incubated in the absence of filipin. In addition, the observation that
only band C CFTR was labeled with a membrane-impermeant biotinylating agent implies that we are detecting only cell surface CFTR.
Characterization of band C CFTR was performed by running a radiolabeled
immunoprecipitate of CFTR from C127-CFTR cells in a parallel lane (data
not shown). Moreover, our inability to detect immature CFTR also
indicates that the concentration of filipin used was not deleterious to cell viability. Caveolae were also disrupted by exposure of the apical
plasma membrane to cholesterol oxidase (0.5 U/ml). This procedure
results in the redistribution of plasma membrane caveolin first to the
endoplasmic reticulum and then to the Golgi complex (41). As with
filipin, data from experiments in which Calu-3 cells were exposed to
cholesterol oxidase showed a loss of plasma membrane caveolae (83 ± 7% by densitometric analysis; Fig.
3A) but no difference in the amount
of cell surface CFTR compared with nontreated cells (Table 2).
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In contrast to the effects of caveolae disrupters, exposure of Calu-3 cells to drugs that disrupt clathrin-coated pathways caused a marked alteration in cell surface CFTR levels. Thus exposure of Calu-3 cells to chlorpromazine (100 µM) resulted in a marked inhibition in the uptake of 125I-transferrin, consistent with a disruption in clathrin-mediated endocytosis (Fig. 3B). When Calu-3 cells were incubated in chlorpromazine for 30 min at 37°C, the level of cell surface CFTR measured in the presence of chlorpromazine was approximately threefold higher than that observed in cells not exposed to chlorpromazine (Fig. 3C, Table 2). These results are not only consistent with a model of CFTR internalization via clathrin-coated vesicles but are also consistent with a model in which CFTR continually recycles between the plasma membrane and an intracellular pool. As with filipin, the observation that only the level of mature band C CFTR was increased in the presence of chlorpromazine (Fig. 3C) indicates that the increased signal is due to cell surface accumulation of CFTR and not the loss of cell integrity. Exposure of cells in the presence and absence of chlorpromazine at 4°C did not result in any significant differences in the ability of Calu-3 cells to bind 125I-transferrin (data not shown). Two other approaches were also utilized to disrupt clathrin-mediated endocytosis, K+ depletion and hypertonic shock. As with chlorpromazine, both K+ depletion and hypertonic shock resulted in a significant inhibition of 125I-transferrin uptake, indicating an ablation of clathrin-coated vesicle formation (Fig. 3B). As with chlorpromazine, exposure of cells to either K+ depletion or hypertonic shock resulted in an accumulation of cell surface CFTR (Table 2).
CFTR but not caveolin-1 can be extracted in Triton
X-100. Caveolae have been isolated based on their
insolubility in Triton X-100 (10, 12, 36). Thus experiments were
performed to determine the solubility of CFTR and caveolin-1 in Triton
X-100. Monolayers of cells were lysed in 1% (vol/vol) Triton X-100
according to the method of Brown and Rose (10) followed by
centrifugation to yield detergent-soluble and detergent-insoluble
fractions. The fractions were subsequently fully solubilized in Laemmli
SDS sample buffer and subjected to immunoblot and immunoprecipitation assays. As expected, caveolin-1 was not detected in the
detergent-soluble pool from Calu-3 cells; however, analysis of
detergent-insoluble domains revealed the presence of caveolin-1 (Fig.
4A).
Similar results were obtained in the C127-CFTR overexpression system. Although some caveolin-1 was present in the detergent-soluble domain,
densitometric analysis showed that this represented only ~5% of the
total caveolin-1 pool. Similar experiments were also performed to
determine the ability of Triton X-100 to solubilize CFTR. Confluent
monolayers of Calu-3 and C127-CFTR cells were separated into
detergent-soluble and -insoluble fractions as described in
Detergent
solubilization. Subsequently, after full
solubilization in RIPA buffer, CFTR was immunoprecipitated with a
monoclonal anti-CFTR antibody. In Calu-3 cells, CFTR was fully
extracted into the detergent-soluble pool (Fig.
4B). No CFTR was observed in the
detergent-insoluble fraction even after prolonged exposure of the gel
to X-ray film. Similar results were obtained from C127-CFTR cells in
which CFTR was also detected in the detergent-soluble fraction.
However, in contrast to Calu-3 cells, CFTR was also detected in the
detergent-insoluble pool of C127-CFTR cells, accounting for
~10-15% of total band C mature CFTR. Because only mature band C
CFTR was observed in the detergent-insoluble pool, we presume that this
reflects CFTR that is present in plasma membrane caveolae and not CFTR
that has aggregated along the biosynthetic pathway as a result of
overexpression (aggregates that are also likely to be insoluble in
Triton X-100).
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CFTR in natively expressing cells is not targeted to
caveolae. Given that the level of caveolin expression
is stoichiometrically related to the number of caveolae, it appears
that Calu-3 cells contain fewer caveolae than C127-CFTR cells. It is
therefore possible that our inability to detect CFTR in Calu-3 cells
(which is also expressed in a lower copy number in Calu-3 cells) in a
detergent-insoluble pool may reflect a limit of detection rather than a
complete absence. To determine whether detergent solubility of CFTR
truly represents an absence of CFTR from caveolae, we isolated caveolae
from Calu-3 and C127-CFTR cells using the method of Smart et al. (42).
Equal amounts of caveolae (10 µg) were resolved by SDS-PAGE.
Antibodies against caveolin-1 showed that similar levels of caveolin-1
were present in the two preparations (Fig.
5A).
Caveolae were also fully solubilized in RIPA buffer and subjected to
immunoprecipitation with CFTR antibodies followed by in vitro
phosphorylation as described in
METHODS. CFTR was detected as a single
band of 180 kDa in caveolae from C127-CFTR cells, consistent with
mature fully glycosylated band C CFTR (Fig.
5B). The observation that only band
C CFTR was detected in caveolae from C127-CFTR cells indicates that
this fraction contains CFTR derived from an apical membrane domain and
is not contaminated with endoplasmic reticulum or Golgi (i.e., nascent
polypeptide chain band A and band B are not detected). In
contrast, no CFTR was detected in caveolae isolated from Calu-3 cells
(Fig. 5B). Our inability to detect
CFTR in caveolae from Calu-3 cells is not due to physicochemical
constraints on the ability of the CFTR antibodies to recognize their
epitope within the confines of caveolae because we can detect CFTR in
caveolae from C127-CFTR cells. In addition, our inability to detect
CFTR in caveolae from Calu-3 cells is not due to differing amounts of
caveolae between Calu-3 and C127-CFTR cells because equal amounts of
caveolae were utilized for immunoprecipitation and detection of CFTR.
Nonetheless, it is possible that there is a small amount of CFTR (below
the resolution limits of our detection system) in caveolae from Calu-3
cells.
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DISCUSSION |
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There is growing evidence that there are at least two major pools of CFTR, one in the apical plasma membrane and one in an early endosomal compartment. Various endocytic pathways have been characterized in terms of morphology, coat composition, and size of vesicle (for a review see Ref. 22). Probably the two best studied and possibly most important pathways are clathrin-coated vesicle-mediated uptake and caveolae-mediated uptake. An immunologic and biophysical study by Bradbury et al. (4) suggested that at least a portion of plasma membrane CFTR enters clathrin-mediated pathways in human cultured epithelial cells and native rat colonic epithelial cells (4). Yet membrane protein internalization via caveolae has also been reported in epithelial cells (21, 30, 40). Thus the aim of our present study was to delineate the pathway(s) for CFTR internalization from the apical plasma membrane.
Our initial experiments were designed to determine whether caveolin is expressed in the same cell type as CFTR. Our studies revealed that although caveolin-1 and caveolin-2 are expressed in some epithelial cells that express CFTR, there are also CFTR-expressing epithelial cells that do not express caveolin. We focused our further studies on Calu-3 cells (a model of human airway serous epithelial cells) and C127-CFTR cells (a stably transfected mouse cell line overexpressing CFTR) because these cell lines express both CFTR and caveolin-1. Detergent insolubility has been associated with the localization of proteins within caveolae (10, 12, 36). Our results show that although CFTR was entirely detergent soluble in the natively expressing cell line Calu-3, significant levels of CFTR were also present in the detergent-insoluble pool of the CFTR-overexpressing cell line C127-CFTR. Although the relative detergent solubility and insolubility of CFTR from Calu-3 and C127-CFTR cells is not definitive proof for the absence of CFTR from caveolae in Calu-3 cells and its presence in caveolae from C127-CFTR cells, the data are consistent with the hypothesis that CFTR is excluded from caveolae but that overexpression of CFTR can result in the appearance of CFTR in caveolae.
The relative roles of clathrin and caveolae-mediated internalization pathways have been investigated for several important membrane transport proteins including GLUT-4 and proton ATPase cells (8, 18, 27, 37). For CFTR to enter caveolae, caveolae must be present within the apical plasma membrane domain of chloride secretory epithelial cells because CFTR is polarized to the apical plasma membrane. Immunocytochemical image analysis reveals that indeed caveolin-1 is present within the apical membrane domain of Calu-3 cells, and because caveolin is a generally accepted marker for caveolae, the potential for CFTR-caveolae interactions in the apical plasma membrane exists. Although there are a few reports (20, 45) indicating that caveolae are polarized to the basolateral plasma membrane in epithelial cells and absent from the apical membrane, there are many reports (13, 24, 25, 36, 43) of caveolae and/or detergent-resistant domains within the apical as well as basolateral domains of polarized epithelial cells.
Several pharmacological tools have been described as differentially affecting clathrin-mediated and caveolae-mediated endocytic pathways. Based on subcellular fractionation data from Bradbury et al. (4), it appears that CFTR is targeted to clathrin-coated pits and hence to clathrin-coated vesicles (4). It has proved very difficultto identify CFTR in clathrin-coated pits and vesicles by immunoelectron microscopy because as an ion channel, CFTR is not expressed at a very high copy number. Chlorpromazine, an amphipathic drug that inhibits clathrin-mediated endocytosis (46), caused a marked reduction in clathrin-mediated pathways in Calu-3 cells as did K+ depletion and hypertonic shock as evidenced by inhibition of 125I-transferrin uptake. Chlorpromazine, K+ depletion, and hypertonic shock also resulted in the accumulation of CFTR at the cell surface, consistent with a model in which CFTR is internalized into early endosomes and recycled back to the cell surface. Our observations are also consistent with those of Lukacs et al. (26), who demonstrated that inhibition of clathrin-mediated endocytosis by hypertonic media prevented delivery of CFTR to early endosomes. Thus although our demonstration of CFTR-clathrin association in the present work is based primarily on the inhibition of clathrin-mediated endocytosis by chlorpromazine, K+ depletion, and hypertonic shock, our present data, together with earlier works by Bradbury et al. (4) and Lukacs et al. (26), provide strong evidence for the internalization of CFTR via clathrin-coated vesicles. Filipin, a sterol binding drug, causes disruption of caveolae and redistribution of caveolae resident proteins (35). Under our experimental conditions, filipin abrogated caveolae as monitored by a loss of caveolin-1 from caveolae. However, in contrast to the effects of chlorpromazine, K+ depletion, and hypertonic shock on CFTR internalization, the level of cell surface CFTR was not altered in the presence of either filipin or cholesterol oxidase. Finally, caveolae isolated from cells that natively express CFTR were devoid of CFTR. However, when overexpressed, it appears that CFTR can enter caveolae.
Many glycosylphosphatidylinositol (GPI)-anchored proteins are found clustered within caveolae. The signal for GPI attachment to proteins consists of a short hydrophobic sequence (13-17 residues) and an appropriate upstream cleavage or attachment site (11). CFTR does not contain such sequences, indicating that at least there is no GPI targeting of CFTR to caveolae, consistent with our inability to detect CFTR in caveolae from cells that natively express CFTR. It is possible that our inability to find CFTR within caveolae represents a limit of our detection abilities. Indeed, even in the CFTR overexpression system C127-CFTR, the level of caveolae CFTR represents only ~10-15% of the total plasma membrane CFTR pool. Thus although it is possible that a small amount of CFTR in natively expressing cells (e.g., Calu-3 cells) is present in caveolae, the physiological significance of these CFTR molecules is likely to be very small. Taken together, our studies support the hypothesis that caveolae do not play a significant role in the trafficking of CFTR in polarized epithelial cells.
These studies extend previous observations by Bradbury et al. (4) defining a role for clathrin-coated vesicles in CFTR trafficking. Indeed, our findings are also confirmed by a recent study (26) showing that perturbation of clathrin-coated pathways inhibits the appearance of CFTR in endosomal compartments in a transformed Chinese hamster ovary cell line expressing CFTR. Such findings are also consistent with other studies suggesting that localization of membrane proteins to either clathrin-coated domains or caveolae is mutually exclusive. Scherer et al. (37) suggested that recombinant GLUT-4, which is internalized through clathrin-coated vesicles, can also enter caveolae. However, the 3T3-L1 system used in these studies may display altered trafficking properties because GLUT-1 and GLUT-4 colocalize in 3T3-L1 cells, whereas they are segregated in tissues (rat adipocytes) that natively express these proteins (18, 27). The observation that CFTR can be found in caveolae of CFTR-overexpressing cells may reflect a "leaking" of CFTR into these domains after insertion into the plasma membrane of high levels of CFTR. Such observations should provide a note of caution when attempting to investigate the cell biology of CFTR in an overexpression system.
In conclusion, we show that CFTR enters pathways for clathrin-mediated internalization and that CFTR appears to be excluded from caveolae. Although we cannot fully exclude the possibility of very low levels of CFTR in caveolae from secretory epithelial cells, if there are any CFTR molecules in caveolae from native epithelia, the physiological consequences of these CFTR molecules are likely to be insignificant.
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ACKNOWLEDGEMENTS |
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We thank Maitrayee Sahu, Lee Ann Giltinan, and Thomas Jackson for excellent technical assistance.
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FOOTNOTES |
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This work was supported by the National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-47850 (to N. A. Bradbury).
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. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: N. A. Bradbury, S306 BST South, Dept. of Cell Biology and Physiology, Univ. of Pittsburgh School of Medicine, 3500 Terrace St., Pittsburgh, PA 15261 (E-mail: nabrad+{at}pitt.edu).
Received 24 August 1998; accepted in final form 23 December 1998.
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